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Principles of Biochemistry

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Principles of Biochemistry

Fifth Edition

Laurence A. Moran

University of Toronto

H. Robert Horton

North Carolina State University

K. Gray Scrimgeour

University of Toronto

Marc D. Perry

University of Toronto

PEARSON

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Library of Congress Cataloging-in-Publication Data

Principles of biochemistry / H. Robert Horton ... [et al]. — 5th ed.
p. cm.

ISBN 0-321-70733-8

1. Biochemistry. I. Horton, H. Robert, 1935-
QP514.2.P745 2012
612'. 015 — dc23

2011019987

ISBN 10: 0-321-70733-8

ISBN 13: 978-0-321-70733-8
123456789 10— DOW— 16 15 14 13 12

PEARSON

www.pearsonhighered.com

Science should be as simple as possible,
but not simpler.

- Albert Einstein

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Brief Contents

Part One

Introduction

1 Introduction to Biochemistry l

2 Water 28

Part Two

Structure and Function

3 Amino Acids and the Primary Structures of Proteins 55

4 Proteins: Three-Dimensional Structure and Function 85

5 Properties of Enzymes 134

6 Mechanisms of Enzymes 162

7 Coenzymes and Vitamins 196

8 Carbohydrates 227

9 Lipids and Membranes 256

Part Three

Metabolism and Bioenergetics

10 Introduction to Metabolism 294

11 Glycolysis 325

12 Gluconeogenesis, the Pentose Phosphate
Pathway, and Glycogen Metabolism 355

13 The Citric Acid Cycle 385

14 Electron Transport and ATP Synthesis 417

15 Photosynthesis 443

16 Lipid Metabolism 475

17 Amino Acid Metabolism 514

18 Nucleotide Metabolism 550

Part Four

Biological Information Flow

19 Nucleic Acids 573

20 DNA Replication, Repair, and Recombination 601

21 Transcription and RNA Processing 634

22 Protein Synthesis 666

Contents

viii

To the Student xxiii

Preface xxv

About the Authors xxxiii

Part One

Introduction

1 Introduction to Biochemistry 1

1.1 Biochemistry Is a Modern Science 2

1.2 The Chemical Elements of Life 3

1.3 Many Important Macromolecules Are Polymers 4

A. Proteins 6

B. Polysaccharides 6

C. Nucleic Acids 7

D. Lipids and Membranes 9

1.4 The Energetics of Life 10

A. Reaction Rates and Equilibria 11

B. Thermodynamics 12

C. Equilibrium Constants and Standard Gibbs Free Energy Changes 13

D. Gibbs Free Energy and Reaction Rates 14

1.5 Biochemistry and Evolution 15

1.6 The Cell Is the Basic Unit of Life 17

1.7 Prokaryotic Cells: Structural Features 17

1.8 Eukaryotic Cells: Structural Features 18

A. The Nucleus 20

B. The Endoplasmic Reticulum and Golgi Apparatus 20

C. Mitochondria and Chloroplasts 21

D. Specialized Vesicles 22

E. The Cytoskeleton 23

1.9 A Picture of the Living Cell 23

1.10 Biochemistry Is Multidisciplinary 26

Appendix: The Special Terminology of Biochemistry 26
Selected Readings 27

2 Water 28

2.1 The Water Molecule Is Polar 29

2.2 Hydrogen Bonding in Water 30
Box 2.1 Extreme Thermophiles 32

2.3 Water Is an Excellent Solvent 32

A. Ionic and Polar Substances Dissolve in Water 32
Box 2.2 Blood Plasma and Seawater 33

B. Cellular Concentrations and Diffusion 34

C. Osmotic Pressure 34

2.4 Nonpolar Substances Are Insoluble in Water 35

CONTENTS ix

2.5 Noncovalent Interactions 37

A. Charge-Charge Interactions 37

B. Hydrogen Bonds 37

C. Van der Waals Forces 38

D. Hydrophobic Interactions 39

2.6 Water Is Nucleophilic 39

Box 2.3 The Concentration of Water 41

2.7 Ionization of Water 41

2.8 The pH Scale 43

Box 2.4 The Little “p” in pH 44

2.9 Acid Dissociation Constants of Weak Acids 44

Sample Calculation 2.1 Calculating the pH of Weak Acid Solutions 49

2.10 Buffered Solutions Resist Changes in pH 50
Sample Calculation 2.2 Buffer Preparation 50
Summary 52

Problems 52
Selected Readings 54

PART TWO

Structure and Function

3 Amino Acids and the Primary Structures of Proteins 55

3.1 General Structure of Amino Acids 56

3.2 Structures of the 20 Common Amino Acids 58

Box 3.1 Fossil Dating by Amino Acid Racemization 58

A. Aliphatic R Groups 59

B. Aromatic R Groups 59

C. R Groups Containing Sulfur 60

D. Side Chains with Alcohol Groups 60
Box 3.2 An Alternative Nomenclature 61

E. Positively Charged R Groups 61

F. Negatively Charged R Groups and Their Amide Derivatives 62

G. The Hydrophobicity of Amino Acid Side Chains 62

3.3 Other Amino Acids and Amino Acid Derivatives 62

3.4 Ionization of Amino Acids 63

Box 3.3 Common Names of Amino Acids 64

3.5 Peptide Bonds Link Amino Acids in Proteins 67

3.6 Protein Purification Techniques 68

3.7 Analytical Techniques 70

3.8 Amino Acid Composition of Proteins 73

3.9 Determining the Sequence of Amino Acid Residues 74

3.10 Protein Sequencing Strategies 76

3.11 Comparisons of the Primary Structures of
Proteins Reveal Evolutionary Relationships 79
Summary 82

Problems 82
Selected Readings 84

4 Proteins: Three-Dimensional Structure and Function 85

4.1 There Are Four Levels of Protein Structure 87

4.2 Methods for Determining Protein Structure 88

X CONTENTS

4.3 The Conformation of the Peptide Group 91

Box 4.1 Flowering Is Controlled by Cis/Trans Switches 93

4.4 The a Helix 94

4.5 (3 Strands and f3 Sheets 97

4.6 Loops and Turns 98

4.7 Tertiary Structure of Proteins 99

A. Supersecondary Structures 100

B. Domains 101

C. Domain Structure, Function, and Evolution 102

D. Intrinsically Disordered Proteins 102

4.8 Quaternary Structure 103

4.9 Protein-Protein Interactions 109

4.10 Protein Denaturation and Renaturation 110

4.11 Protein Folding and Stability 114

A. The Hydrophobic Effect 114

B. Hydrogen Bonding 115

Box 4.2 CASP: The Protein Folding Game 116

C. Van der Waals Interactions and Charge-Charge Interactions 117

D. Protein Folding Is Assisted by Molecular Chaperones 117

4.12 Collagen, a Fibrous Protein 119
Box 4.3 Stronger Than Steel 121

4.13 Structure of Myoglobin and Hemoglobin 122

4.14 Oxygen Binding to Myoglobin and Hemoglobin 123

A. Oxygen Binds Reversibly to Heme 123

B. Oxygen-Binding Curves of Myoglobin and Hemoglobin 124
Box 4.4 Embryonic and Fetal Hemoglobins 126

C. Hemoglobin Is an Allosteric Protein 127

4.15 Antibodies Bind Specific Antigens 129
Summary 130

Problems 131
Selected Readings 133

5 Properties of Enzymes 134

5.1 The Six Classes of Enzymes 136

Box 5.1 Enzyme Classification Numbers 137

5.2 Kinetic Experiments Reveal Enzyme Properties 138

A. Chemical Kinetics 138

B. Enzyme Kinetics 139

5.3 The Michaelis-Menten Equation 140

A. Derivation of the Michaelis-Menten Equation 141

B. The Calalytic Constant K cat 143

C. The Meanings of K m 144

5.4 Kinetic Constants Indicate Enzyme Activity and Catalytic Proficiency

5.5 Measurement of K m and l/ max 145

Box 5.2 Hyperbolas Versus Straight Lines 146

5.6 Kinetics of Multisubstrate Reactions 147

5.7 Reversible Enzyme Inhibition 148

A. Competitive Inhibition 149

B. Uncompetitive Inhibition 150

144

CONTENTS Xi

C. Noncompetitive Inhibition 150

D. Uses of Enzyme Inhibition 151

5.8 Irreversible Enzyme Inhibition 152

5.9 Regulation of Enzyme Activity 153

A. Phosphofructokinase Is an Allosteric Enzyme 154

B. General Properties of Allosteric Enzymes 155

C. Two Theories of Allosteric Regulation 156

D. Regulation by Covalent Modification 158

5.10 Multienzyme Complexes and Multifunctional Enzymes 158
Summary 159

Problems 159
Selected Readings 161

6 Mechanisms of Enzymes 162

6.1 The Terminology of Mechanistic Chemistry 162

A. Nucleophilic Substitutions 163

B. Cleavage Reactions 163

C. Oxidation-Reduction Reactions 164

6.2 Catalysts Stabilize Transition States 164

6.3 Chemical Modes of Enzymatic Catalysis 166

A. Polar Amino Acids Residues in Active Sites 166

Box 6.1 Site-Directed Mutagenesis Modifies Enzymes 167

B. Acid-Base Catalysis 168

C. Covalent Catalysis 169

D. pH Affects Enzymatic Rates 170

6.4 Diffusion-Controlled Reactions 171

A. Triose Phosphate Isomerase 172

Box 6.2 The “Perfect Enzyme”? 174

B. Superoxide Dismutase 175

6.5 Modes of Enzymatic Catalysis 175

A. The Proximity Effect 176

B. Weak Binding of Substrates to Enzymes 178

C. Induced Fit 179

D. Transition State Stabilization 180

6.6 Serine Proteases 183

A. Zymogens Are Inactive Enzyme Precursors 183
Box 6.3 Kornberg’s Ten Commandments 183

B. Substrate Specificity of Serine Proteases 184

C. Serine Proteases Use Both the Chemical
and the Binding Modes of Catalysis 185

Box 6.4 Clean Clothes 186
Box 6.5 Convergent Evolution 187

6.7 Lysozyme 187

6.8 Arginine Kinase 190
Summary 192
Problems 193
Selected Readings 194

His-95

Xii CONTENTS

Coenzymes and Vitamins 196

7.1 Many Enzymes Require Inorganic Cations 197

7.2 Coenzyme Classification 197

7.3 ATP and Other Nucleotide Cosubstrates 198
Box 7.1 Missing Vitamins 200

7.4 NAD© and NADP© 200

Box 7.2 NAD Binding to Dehydrogenases 203

7.5 FAD and FMN 204

7.6 Coenzyme A and Acyl Carrier Protein 204

7.7 Thiamine Diphosphate 206

7.8 Pyridoxal Phosphate 207

7.9 Vitamin C 209

7.10 Biotin 211

Box 7.3 One Gene: One Enzyme 212

7.11 Tetrahydrofolate 213

7.12 Cobalamin 215

7.13 Lipoamide 216

7.14 Lipid Vitamins 217

A. Vitamin A 217

B. Vitamin D 218

C. Vitamin E 218

D. Vitamin K 218

7.15 Ubiquinone 219

Box 7.4 Rat Poison 220

7.16 Protein Coenzymes 221

7.17 Cytochromes 221

Box 7.5 Noble Prizes for Vitamins and Coenzymes 223
Summary 223
Problems 224
Selected Readings 226

8 Carbohydrates 227

8.1 Most Monosaccharides Are Chiral Compounds 228

8.2 Cyclization of Aldoses and Ketoses 230

8.3 Conformations of Monosaccharides 234

8.4 Derivatives of Monosaccharides 235

A. Sugar Phosphates 235

B. Deoxy Sugars 235

C. Amino Sugars 235

D. Sugar Alcohols 236

E. Sugar Acids 236

8.5 Disaccharides and Other Glycosides 236

A. Structures of Disaccharides 237

B. Reducing and Nonreducing Sugars 238

C. Nucleosides and Other Glycosides 239
Box 8.1 The Problem with Cats 240

8.6 Polysaccharides 240

A. Starch and Glycogen 240

B. Cellulose 243

CONTENTS Xiii

C. Chitin 244

8.7 Glycoconjugates 244

A. Proteoglycans 244

Box 8.2 Nodulation Factors Are Lipo-Oligosaccharides 246

B. Peptidoglycans 246

C. Glycoproteins 248

Box 8.3 ABO Blood Group 250
Summary 252
Problems 253
Selected Readings 254

9 Lipids and Membranes 256

9.1 Structural and Functional Diversity of Lipids 256

9.2 Fatty Acids 256

Box 9.1 Common Names of Fatty Acids 258
Box 9.2 Trans Fatty Acids and Margarine 259

9.3 Triacylglycerols 261

9.4 Glycerophospholipids 262

9.5 Sphingolipids 263

9.6 Steroids 266

9.7 Other Biologically Important Lipids 268

9.8 Biological Membranes 269

A. Lipid Bilayers 269

Box 9.3 Gregor Mendel and Gibberellins 270

B. Three Classes of Membrane Proteins 270
Box 9.4 New Lipid Vesicles, or Liposomes 272

Box 9.5 Some Species Have Unusual Lipids in Their Membranes 274

C. The Fluid Mosaic Model of Biological Membranes 274

9.9 Membranes Are Dynamic Structures 275

9.10 Membrane Transport 277

A. Thermodynamics of Membrane Transport 278

B. Pores and Channels 279

C. Passive Transport and Facilitated Diffusion 280

D. Active Transport 282

E. Endocytosis and Exocytosis 283

9.11 Transduction of Extracellular Signals 283

A. Receptors 283

Box 9.6 The Hot Spice of Chili Peppers 284

B. Signal Transducers 285

C. The Adenylyl Cyclase Signaling Pathway 287

D. The Inositol-Phospholipid Signaling Pathway 287
Box 9.7 Bacterial Toxins and G Proteins 290

E. Receptor Tyrosine Kinases 290
Summary 291

Problems 292
Selected Readings 293

Xiv CONTENTS

PART THREE

Metabolism and Bioenergetics

10 Introduction to Metabolism 294

10.1 Metabolism Is a Network of Reactions 294

10.2 Metabolic Pathways 297

A. Pathways Are Sequences of Reactions 297

B. Metabolism Proceeds by Discrete Steps 297

C. Metabolic Pathways Are Regulated 297

D. Evolution of Metabolic Pathways 301

10.3 Major Pathways in Cells 302

10.4 Compartmentation and Interorgan Metabolism 304

10.5 Actual Gibbs Free Energy Change, Not Standard Free Energy Change,
Determines the Direction of Metabolic Reactions 306

Sample Calculation 10.1 Calculating Standard Gibbs Free Energy
Change from Energies of Formation 308

10.6 The Free Energy of ATP Hydrolysis 308

10.7 The Metabolic Roles of ATP 311

A. Phosphoryl Group Transfer 311

Sample Calculation 10.2 Gibbs Free Energy Change 312
Box 10.1 The Squiggle 312

B. Production of ATP by Phosphoryl Group Transfer 314

C. Nucleotidyl Group Transfer 315

10.8 Thioesters Have High Free Energies of Hydrolysis 316

10.9 Reduced Coenzymes Conserve Energy from Biological Oxidations 316

A. Gibbs Free Energy Change Is Related to Reduction Potential 317

B. Electron Transfer from NADH Provides Free Energy 319

Box 10.2 NAD© and NADH Differ in Their Ultraviolet Absorption Spectra

10.10 Experimental Methods for Studying Metabolism 321
Summary 322

Problems 323
Selected Readings 324

1 1 Glycolysis 325

11.1 The Enzymatic Reactions of Glycolysis 326

11.2 The Ten Steps of Glycolysis 326

1. Hexokinase 326

2. Glucose 6-Phosphate Isomerase 327

3. Phosphofructokinase-1 330

4. Aldolase 330

Box 11.1 A Brief History of the Glycolysis Pathway 331

5. Triose Phosphate Isomerase 332

6. Glyceraldehyde 3-Phosphate Dehydrogenase 333

7. Phosphoglycerate Kinase 335

Box 11.2 Formation of 2,3-S/sphosphoglycerate in Red Blood Cells 335
Box 11.3 Arsenate Poisoning 336

8. Phosphoglycerate Mutase 336

9. Enolase 338

lO.Pryuvate Kinase 338

321

CONTENTS XV

11.3 The Fate of Pryuvate 338

A. Metabolism of Pryuvate to Ethanol 339

B. Reduction of Pyruvate to Lactate 340

Box 11.4 The Lactate of the Long-Distance Runner 341

11.4 Free Energy Changes in Glycolysis 341

11.5 Regulation of Glycolysis 343

A. Regulation of Hexose Transporters 344

B. Regulation of Hexokinase 344

Box 11.5 Glucose 6-Phosphate Has a Pivotal Metabolic Role in the Liver 345

C. Regulation of Phosphofructokinase-1 345

D. Regulation of Pyruvate Kinase 346

E. The Pasteur Effect 347

11.6 Other Sugars Can Enter Glycolysis 347

A. Sucrose Is Cleaved to Monosaccharides 348

B. Fructose Is Converted to Glyceraldehyde 3-Phosphate 348

C. Galactose Is Converted to Glucose 1-Phosphate 349
Box 11.6 A Secret Ingredient 349

D. Mannose Is Converted to Fructose 6-Phosphate 351

11.7 The Entner-Doudoroff Pathway in Bacteria 351
Summary 352

Problems 353
Selected Readings 354

12 Gluconeogenesis, the Pentose Phosphate Pathway,
and Glycogen Metabolism 355

12.1 Gluconeogenesis 356

A. Pyruvate Carboxylase 357

B. Phosphoenolpyruvate Carboxykinase 358

C. Fructose 1,6-b/sphosphatase 358
Box 12.1 Supermouse 359

D. Glucose 6-Phosphatase 359

12.2 Precursors for Gluconeogenesis 360

A. Lactate 360

B. Amino Acids 360

C. Glycerol 361

D. Propionate and Lactate 361

E. Acetate 362

Box 12.2 Glucose Is Sometimes Converted to Sorbitol 362

12.3 Regulation of Gluconeogenesis 363

Box 12.3 The Evolution of a Complex Enzyme 364

12.4 The Pentose Phosphate Pathway 364

A. Oxidative Stage 366

B. Nonoxidative Stage 364

Box 12.4 Glucose 6-Phosphate Dehydrogenase Deficiency in Humans 367

C. Interconversions Catalyzed by Transketolase and Transaldolase 368

12.5 Glycogen Metabolism 368

A. Glycogen Synthesis 369

B. Glycogen Degradation 370

12.6 Regulation of Glycogen Metabolism in Mammals 372

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XVi CONTENTS

A. Regulation of Glycogen Phosphorylase 372
Box 12.5 Head Growth and Tail Growth 373

B. Hormones Regulate Glycogen Metabolism 375

C. Hormones Regulate Gluconeogenesis and Glycolysis 376

12.7 Maintenance of Glucose Levels in Mammals 378

12.8 Glycogen Storage Diseases 381
Summary 382

Problems 382
Selected Readings 383

1 3 The Citric Acid Cycle 385

Box 13.1 An Egregious Error 386

13.1 Conversion of Pyruvate to Acetyl CoA 387

Sample Calculation 13.1 390

13.2 The Citric Acid Cycle Oxidizes Acetyl CoA 391
Box 13.2 Where Do the Electrons Come From? 392

13.3 The Citric Acid Cycle Enzymes 394

1. Citrate Synthase 394

Box 13.3 Citric Acid 396

2. Aconitase 396

Box 13.4 Three-Point Attachment of Prochiral Substrates to Enzymes

3. Isocitrate Dehydrogenase 397

4. The a-Ketoglutarate Dehydrogenase Complex 398

5. Succinyl CoA Synthetase 398

6. Succinate Dehydrogenase Complex 399
Box 13.5 What’s in a Name? 399

Box 13.6 On the Accuracy of the World Wide Web 401

7 . Fumarase 401

8. Malate Deydrogenase 401

Box 13.7 Converting One Enzyme into Another 402

13.4 Entry of Pyruvate Into Mitochondria 402

13.5 Reduced Coenzymes Can Fuel the Production of ATP 405

13.6 Regulation of the Citric Acid Cycle 406

13.7 The Citric Acid Cycle Isn’t Always a "Cycle” 407
Box 13.8 A Cheap Cancer Drug? 408

13.8 The Glyoxylate Pathway 409

13.9 Evolution of the Citric Acid Cycle 412
Summary 414

Problems 414
Selected Readings 416

14 Electron Transport and ATP Synthesis 417

14.1 Overview of Membrane-associated Electron Transport

and ATP Synthesis 418

14.2 The Mitochondrion 418

Box 14.1 An Exception to Every Rule 420

14.3 The Chemiosmotic Theory and the Protonmotive Force 420

A. Historical Background: The Chemiosmotic Theory 420

B. The Protonmotive Force 421

397

CONTENTS

XVII

14.4 Electron Transport 423

A. Complexes I Through IV 423

B. Cofactors in Electron Transport 425

14.5 Complex I 426

14.6 Complex II 427

14.7 Complex III 428

14.8 Complex IV 431

14.9 Complex V: ATP Synthase 433

Box 14.2 Proton Leaks and Heat Production 435

14.10 Active Transport of ATP, ADP, and Pj Across
the Mitochondrial Membrane 435

14.11 The P/O Ratio 436

14.12 NADH Shuttle Mechanisms in Eukaryotes 436
Box 14.3 The High Cost of Living 439

14.13 Other Terminal Electron Acceptors and Donors 439

14.14 Superoxide Anions 440
Summary 441
Problems 441
Selected Readings 442

1 5 Photosynthesis 443

15.1 Light-Gathering Pigments 444

A. The Structures of Chlorophylls 444

B. Light Energy 445

C. The Special Pair and Antenna Chlorophylls 446
Box 15.1 Mendel’s Seed Color Mutant 447

D. Accessory Pigments 447

15.2 Bacterial Photosystems 448

A. Photosystem II 448

B. Photosystem I 450

C. Coupled Photosystems and Cytochrome bf 453

D. Reduction Potentials and Gibbs Free Energy in Photosynthesis 455

E. Photosynthesis Takes Place Within Internal Membranes 457
Box 15.2 Oxygen “Pollution” of Earth’s Atmosphere 457

15.3 Plant Photosynthesis 458

A. Chloroplasts 458

B. Plant Photosystems 459

C. Organization of Cloroplast Photosystems 459
Box 15.3 Bacteriorhodopsin 461

15.4 Fixation of C0 2 : The Calvin Cycle 461

A. The Calvin Cycle 462

B. Rubisco: R ibu lose 1,5-b/sphosphate Carboxylase-oxygenase 462

C. Oxygenation of Ribulose 1,5-b/sphosphate 465
Box 15.4 Building a Better Rubisco 466

D. Calvin Cycle: Reduction and Regeneration Stages 466

15.5 Sucrose and Starch Metabolism in Plants 467
Box 15.5 Gregor Mendel’s Wrinkled Peas 469

15.6 Additional Carbon Fixation Pathways 469
A. Compartmentalization in Bacteria 469

XViii CONTENTS

B. The C 4 Pathway 469

C. Crassulacean Acid Metabolism (CAM) 471
Summary 472

Problems 473
Selected Readings 474

16 Lipid Metabolism 475

16.1 Fatty Acid Synthesis 475

A. Synthesis of Malonyl ACP and Acetyl ACP 476

B. The Initiation Reaction of Fatty Acid Synthesis 477

C. The Elongation Reactions of Fatty Acid Synthesis 477

D. Activation of Fatty Acids 479

E. Fatty Acid Extension and Desaturation 479

16.2 Synthesis of Triacylglycerols and Glycerophospholipids 481

16.3 Synthesis of Eicosanoids 483

Box 16.1 s/7-G lycerol 3-Phosphate 484

Box 16.2 The Search for a Replacement for Asprin 486

16.4 Synthesis of Ether Lipids 487

16.5 Synthesis of Sphingolipids 488

16.6 Synthesis of Cholesterol 488

A. Stage 1: Acetyl CoA to Isopentenyl Diphosphate 488

B. Stage 2: Isopentenyl Diphosphate to Squalene 488

C. Stage 3: Squalene to Cholesterol 490

D. Other Products of Isoprenoid Metabolism 490
Box 16.3 Lysosomal Storage Diseases 492

Box 16.4 Regulating Cholesterol Levels 493

16.7 Fatty Acid Oxidation 494

A. Activation of Fatty Acids 494

B. The Reactions of p-Oxidation 494

C. Fatty Acid Synthesis and p-Oxidation 497

D. Transport of Fatty Acyl CoA into Mitochondria 497
Box 16.5 A Trifunctional Enzyme for p-Oxidation 498

E. ATP Generation from Fatty Acid Oxidation 498

F. p-Oxidation of Odd-Chain and Unsaturated Fatty Acids 499

16.8 Eukaryotic Lipids Are Made at a Variety of Sites 501

16.9 Lipid Metabolism Is Regulated by Hormones in Mammals 502

16.10 Absorption and Mobilization of Fuel Lipids in Mammals 505

A. Absorption of Dietary Lipids 505

B. Lipoproteins 505

Box 16.6 Extra Virgin Olive Oil 506

Box 16.7 Lipoprotein Lipase and Coronary Heart Disease 507

C. Serum Albumin 508

16.11 Ketone Bodies Are Fuel Molecules 508

A. Ketone Bodies Are Synthesized in the Liver 509

B. Ketone Bodies Are Oxidized in Mitochondria 510
Box 16.8 Lipid Metabolism in Diabetes 511
Summary 511

Problems 511
Selected Readings 513

CONTENTS xix

17 Amino Acid Metabolism 514

17.1 The Nitrogen Cycle and Nitrogen Fixation 515

17.2 Assimilation of Ammonia 518

A. Ammonia Is Incorporated into Glutamate and Glutamine 518

B. Transamination Reactions 518

17.3 Synthesis of Amino Acids 520

A. Aspartate and Asparagine 520

B. Lysine, Methionine, Threonine 520

C. Alanine, Valine, Leucine, and Isoleucine 521

Box 17.1 Childhood Acute Lymphoblastic Leukemia Can Be Treated
with Asparaginase 522

D. Glutamate, Glutamine, Arginine, and Proline 523

E. Serine, Glycine, and Cysteine 523

F. Phenylalanine, Tyrosine, and Tryptophan 523

G. Histidine 527

Box 17.2 Genetically Modified Food 528

Box 17.3 Essential and Nonessential Amino Acids in Animals 529

17.4 Amino Acids as Metabolic Precursors 529

A. Products Derived from Glutamate, Glutamine, and Aspartate 529

B. Products Derived from Serine and Glycine 529

C. Synthesis of Nitric Oxide from Arginine 530

D. Synthesis of Lignin from Phenylalanine 531

E. Melanin Is Made from Tyrosine 531

17.5 Protein Turnover 531

Box 17.4 Apoptosis-Programmed Cell Death 534

17.6 Amino Acid Catabolism 534

A. Alanine, Asparagine, Aspartate, Glutamate, and Glutamine 535

B. Arginine, Histidine, and Proline 535

C. Glycine and Serine 536

D. Threonine 537

E. The Branched Chain Amino Acids 537

F. Methionine 539

Box 17.5 Phenylketonuria, a Defect in Tyrosine Formation 540

G. Cysteine 540

H. Phenylalanine, Tryptophane, and Tyrosine 541

I. Lysine 542

17.7 The Urea Cycle Converts Ammonia into Urea 542

A. Synthesis of Carbamoyl Phosphate 543

B. The Reactions of the Urea Cycle 543

Box 17.6 Diseases of Amino Acid Metabolism 544

C. Ancillary Reactions of the Urea Cycle 547

17.8 Renal Glutamine Metabolism Produces Bicarbonate 547
Summary 548

Problems 548
Selected Readings 549

18 Nucleotide Metabolism 550

18.1 Synthesis of Purine Nucleotides 550

Box 18.1 Common Names of the Bases 552

18.2 Other Purine Nucleotides Are Synthesized from IMP 554

18.3 Synthesis of Pyrimidine Nucleotides 555

XX CONTENTS

A. The Pathway for Pyrimidine Synthesis 556

Box 18.2 How Some Enzymes Transfer Ammonia from Glutamate 558

B. Regulation of Pyrimidine Synthesis 559

18.4 CTP Is Synthesized from UMP 559

18.5 Reduction of Ribonucleotides to Deoxyribonucleotides 560

18.6 Methylation of dUMP Produces dTMP 560

Box 18.3 Free Radicals in the Reduction of Ribonucleotides 562
Box 18.4 Cancer Drugs Inhibit dTTP Synthesis 564

18.7 Modified Nucleotides 564

18.8 Salvage of Purines and Pyrimidines 564

18.9 Purine Catabolism 565

18.10 Pyrimidine Catabolism 568

Box 18.5 Lesch-Nyhan Syndrome and Gout 569
Summary 571
Problems 571
Selected Readings 572

PART FOUR

Biological Information Flow

19 Nucleic Acids 573

19.1 Nucleotides Are the Building Blocks of Nucleic Acids 574

A. Ribose and Deoxyribose 574

B. Purines and Pyrimidines 574

C. Nucleosides 575

D. Nucleotides 577

19.2 DNA Is Double-Stranded 579

A. Nucleotides Are Joined by 3'-5' Phosphodiester Linkages 580

B. Two Antiparallel Strands Form a Double Helix 581

C. Weak Forces Stabilize the Double Helix 583

D. Conformations of Double-Stranded DNA 585

19.3 DNA Can Be Supercoiled 586

19.4 Cells Contain Several Kinds of RNA 587
Box 19.1 Pulling DNA 588

19.5 Nucleosomes and Chromatin 588

A. Nucleosomes 588

B. Higher Levels of Chromatin Structure 590

C. Bacterial DNA Packaging 590

19.6 Nucleases and Hydrolysis of Nucleic Acids 591

A. Alkaline Hydrolysis of RNA 591

B. Hydrolysis of RNA by Ribonuclease A 592

C. Restriction Endonucleases 593

D. EcoR\ Binds Tightly to DNA 595

19.7 Uses of Restriction Endocucleases 596

A. Restriction Maps 596

B. DNA Fingerprints 596

C. Recombinant DNA 597
Summary 598
Problems 599

Selected Readings 599

CONTENTS XXi

20 DNA Replication, Repair, and Recombination 601

20.1 Chromosomal DNA Replication Is Bidirectional 602

20.2 DNA Polymerase 603

A. Chain Elongation Isa Nucleotidyl-Group-Transfer Reaction 604

B. DNA Polymerase III Remains Bound to the Replication Fork 606

C. Proofreading Corrects Polymerization Errors 607

20.3 DNA Polymerase Synthesizes Two Strands Simultaneously 607

A. Lagging Strand Synthesis Is Discontinuous 608

B. Each Okazaki Fragment Begins with an RNA Primer 608

C. Okazaki Fragments Are Joined by the Action of DNA Polymerase I
and DNA Ligase 609

20.4 Model of the Replisome 610

20.5 Initiation and Termination of DNA Replication 615

20.6 DNA Replication in Eukaryotes 615

A. The Polymerase Chain Reaction Uses DNA Polymerase to
Amplify Selected DNA Sequences 615

B. Sequencing DNA Using Dideoxynucleotides 616

C. Massively Parallel DNA Sequencing by Synthesis 618

20.7 DNA Replication in Eukaryotes 619

20.8 Repair of Damaged DNA 622

A. Repair after Photodimerization: An Example of Direct Repair 622

B. Excision Repair 624

BOX 20.1 The Problem with Methylcytosine 626

20.9 Homologous Recombination 626

A. The Holliday Model of General Recombination 626

B. Recombination in E. coli 627

BOX 20.2 Molecular Links Between DNA Repair and Breast Cancer 630

C. Recombination Can Be a Form of Repair 631
Summary 631

Problems 632
Selected Readings 632

21 Transcription and RNA Processing 633

21.1 Types of RNA 634

21.2 RNA Polymerase 635

A. RNA Polymerase Is an Oligomeric Protein 635

B. The Chain Elongation Reaction 636

21.3 Transcription Initiation 638

A. Genes Have a 5'^3' Orientation 638

B. The Transcription Complex Assembles at a Promoter 639

C. The a sigma Subunit Recognizes the Promoter 640

D. RNA Polymerase Changes Conformation 641

21.4 Transcription Termination 643

21.5 Transcription in Eukaryotes 645

A. Eukaryotic RNA Polymerases 645

B. Eukaryotic Transcription Factors 647

C. The Role of Chromatin in Eukaryotic Transcription 648

21.6 Transcription of Genes Is Regulated 648

21.7 The lac Operon, an Example of Negative and Positive Regulation 650

A. lac Repressor Blocks Transcription 650

B. The Structure of lac Repressor 651

XXii CONTENTS

C. cAMP Regulatory Protein Activates Transcription 652

21.8 Post-transcriptional Modification of RNA 654

A. Transfer RNA Processing 654

B. Ribosomal RNA Processing 655

21.9 Eukaryotic mRNA Processing 655

A. Eukaryotic mRNA Molecules Have Modified Ends 657

B. Some Eukaryotic mRNA Precursors Are Spliced 657
Summary 663

Problems 663
Selected Readings 664

22 Protein Synthesis 665

22.1 The Genetic Code 665

22.2 Transfer RNA 668

A. The Three-Dimensional Structure of tRNA 668

B. tRNA Anticodons Base-Pair with mRNA Codons 669

22.3 Aminoacyl-tRNA Synthetases 670

A. The Aminoacyl-tRNA Synthetase Reaction 671

B. Specificity of Aminoacyl-tRNA Synthetases 671

C. Proofreading Activity of Aminoacyl-tRNA Synthetases 673

22.4 Ribosomes 673

A. Ribosomes Are Composed of Both Ribosomal RNA and Protein 674

B. Ribosomes Contain Two Aminoacyl-tRNA Binding Sites 675

22.5 Initiation of Translation 675

A. Initiator tRNA 675

B. Initiation Complexes Assemble Only at Initiation Codons 676

C. Initiation Factors Help Form the Initiation Complex 677

D. Translation Initiation in Eukaryotes 679

22.6 Chain Elongation During Protein Synthesis Is a Three-Step Microcycle 679

A. Elongation Factors Dock an Aminoacyl-tRNA in the A Site 680

B. Peptidyl Transferase Catalyzes Peptide Bond Formation 681

C. Translocation Moves the Ribosome by One Codon 682

22.7 Termination of Translation 684

22.8 Protein Synthesis Is Energetically Expensive 684

22.9 Regulation of Protein Synthesis 685

A. Ribosomal Protein Synthesis Is Coupled to Ribosome
Assembly in E. coli 685

Box 22.1 Some Antibiotics Inhibit Protein Synthesis 686

B. Globin Synthesis Depends on Heme Availability 687

C. The E. coli trp Operon Is Regulated by Repression and Attenuation 687

22.10 Post-translational Processing 689

A. The Signal Hypothesis 691

B. Glycosylation of Proteins 694
Summary 694

Problems 695
Selected Readings 696
Solutions 697
Glossary 751
Illustration Credits 767
Index 769

To the Student

Welcome to biochemistry — the study of life at the molecular level. As you venture into
this exciting and dynamic discipline, you’ll discover many new and wonderful things.
You’ll learn how some enzymes can catalyze chemical reactions at speeds close to theo-
retical limits — reactions that would otherwise occur only at imperceptibly low rates.
You’ll learn about the forces that maintain biomolecular structure and how even some
of the weakest of those forces make life possible. You’ll also learn how biochemistry has
thousands of applications in day-to-day life — in medicine, drug design, nutrition,
forensic science, agriculture, and manufacturing. In short, you’ll begin a journey of dis-
covery about how biochemistry makes life both possible and better.

Before we begin, we would like to offer a few words of advice:

Don’t just memorize facts; instead, understand principles

In this book, we have tried to identify the most important principles of biochemistry.
Because the knowledge base of biochemistry is continuously expanding, we must grasp
the underlying themes of this science in order to understand it. This textbook is de-
signed to expand on the foundation you have acquired in your chemistry and biology
courses and to provide you with a biochemical framework that will allow you to under-
stand new phenomena as you meet them.

Be prepared to learn a new vocabulary

An understanding of biochemical facts requires that you learn a biochemical vocabu-
lary. This vocabulary includes the chemical structures of a number of key molecules.
These molecules are grouped into families based on their structures and functions. You
will also learn how to distinguish among members of each family and how small mole-
cules combine to form macromolecules such as proteins and nucleic acids.

Test your understanding

True mastery of biochemistry lies with learning how to apply your knowledge and how
to solve problems. Each chapter concludes with a set of carefully crafted problems that
test your understanding of core principles. Many of these problems are mini case stud-
ies that present the problem within the context of a real biochemical puzzle.

For more practice, we are pleased to refer you to The Study Guide for Principles of
Biochemistry by Scott Lefler and Allen Seism which presents a variety of supplementary
questions that you may find helpful. You will also find additional problems on
TheChemistryPlace® for Principles of Biochemistry (http://www.chemplace.com).

Learn to visualize in 3-D

Biochemicals are three-dimensional objects. Understanding what happens in a bio-
chemical reaction at the molecular level requires that you be able to “see” what happens
in three dimensions. We present the structures of simple molecules in several different
ways in order to illustrate their three-dimensional conformation. In addition to the art
in the book, you will find many animations and interactive molecular models on the
website. We strongly suggest you look at these movies and do the exercises that accom-
pany them as well as participate in the molecular visualization tutorials.

Feedback

Finally, please let us know of any errors or omissions you encounter as you use this text.
Tell us what you would like to see in the next edition. With your help we will continue to
evolve this work into an even more useful tool. Our e-mail addresses are at the end of
the Preface. Good luck, and enjoy!

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Preface

Given the breadth of coverage and diversity of ways to present topics in biochemistry,
we have tried to make the text as modular as possible to allow for greater flexibility and
organization. Each large topic resides in its own section. Reaction mechanisms are often
separated from the main thread of the text and can be passed over by those who prefer
not to cover this level of detail. The text is extensively cross-referenced to make it easier
for you to reorganize the chapters and for students to see the interrelationships among
various topics and to drill down to deeper levels of understanding.

We built the book explicitly for the beginning student taking a first course in bio-
chemistry with the aim of encouraging students to think critically and to appreciate
scientific knowledge for its own sake. Parts One and Two lay a solid foundation of
chemical knowledge that will help students understand, rather than merely memo-
rize, the dynamics of metabolic and genetic processes. These sections assume that stu-
dents have taken prerequisite courses in general and organic chemistry and have ac-
quired a rudimentary knowledge of the organic chemistry of carboxylic acids,
amines, alcohols, and aldehydes. Even so, key functional groups and chemical proper-
ties of each type of biomolecule are carefully explained as their structures and func-
tions are presented.

We also assume that students have previously taken a course in biology where they
have learned about evolution, cell biology, genetics, and the diversity of life on this
planet. We offer brief refreshers on these topics wherever possible.

New to this Edition

We are grateful for all the input we received on the first four editions of this text. You’ll
notice the following improvements in this fifth edition:

• Key Concept margin notes are provided throughout to highlight key concepts and
principles that students must know.

• Interest Boxes have been updated and expanded, with 45% new to the fifth edition.
We use interest boxes to explain some topics in more detail, to illustrate certain prin-
ciples with specific examples, to stimulate students curiosity about science, to show
applications of biochemistry, and to explain clinical relevance. We have also added a
few interests boxes that warn students about misunderstanding and misapplications
of biochemistry. Examples include Blood Plasma and Sea Water; Fossil Dating by
Amino Acid Racemization; Embryonic and Fetal Hemoglobins; Clean Clothes; The
Perfect Enzyme; Supermouse; The Evolution of a Complex Enzyme; An Egregious
Error; Mendels Seed Color Mutant; Oxygen Pollution of Earth’s Atmosphere; Extra
Virgin Olive Oil; Missing Vitamins; Pulling DNA; and much more.

• New Material has been added throughout, including an improved explanation of
early evolution (the Web of Life), more emphasis on protein protein interactions, a
new section on intrinsically disordered proteins, and a better description of the dis-
tinction between Gibbs free energy changes and reaction rates. We have removed
the final chapter on Recombinant DNA Technology and integrated much of that
material into earlier chapters. We have added descriptions of a number of new pro-
tein structures and integrated them into two major themes: structure- function and
multienzyme complexes. The best example is the fatty acid synthase complex in
Chapter 16.

In some cases new material was necessary because recent discoveries have
changed our view of some reactions and processes. We now know, for example, that
older versions of uric acid catabolism were incorrect, the correct pathway is shown in
Figure 18.23.

XXV

We have been careful not to add extra detail unless it supports and extends the
basic concepts and principles that we have established over the past four editions.
Similarly, we do not introduce new subjects unless they illustrate new concepts that
were not covered in previous editions. The goal is to keep this textbook focused on
the fundamentals that students need to know and prevent it from bloating up into
an encyclopedia of mostly irrelevant information that detracts from the main
pedagogical goals.

• Selected Readings after each chapter reflect the most current literature and these
have been updated and extended where necessary. We have added over 120 new
references and deleted many that are no longer appropriate. Although we have al-
ways included references to the pedagogical literature, you will note that we have
added quite a few more references of this type. Students now have easy access to
these papers and they are often more informative than advanced papers in the
purely scientific literature.

• Art is an important component of a good textbook. Our art program has been ex-
tensively revised, with many new photos to illustrate concepts explained in the text;
new and updated ribbon art, and improved versions of many figures. Many of the
new photos are designed to attract and/or hold the students attention. They can be
powerful memory aids and some of them are used to lighten up the subject in a
way that is rarely seen in other textbooks (see page 204). We believe that the look
and feel of the book has been much improved, making it more appealing to stu-
dents without sacrificing any of the rigor and accuracy that has been a hallmark of
previous editions.

A focus on principles

There are, in essence, two kinds of biochemistry textbooks: those for reference and
those for teaching. It is difficult for one book to be both as it is those same thickets of
detail sought by the professional that ensnare the struggling novice on his or her first
trip through the forest. This text is unapologetically a text for teaching. It has been de-
signed to foster student understanding and is not an encyclopedia of biochemistry. This
book focuses unwaveringly on teaching basic principles and concepts, each principle
supported by carefully chosen examples. We really do try to get students to see the forest
and not the trees!

Because of this focus, the material in this book can be covered in a two-semester
course without having to tell students to skip certain chapters or certain sections. The
book is also suitable for a one-semester course that concentrates on certain aspects of
biochemistry where some subjects are not covered. Instructors can be confident that the
core principles and concepts are explained thoroughly and correctly.

A focus on chemistry

When we first wrote this text, we decided to take the time to explain in chemical terms
the principles that we want to emphasize. In fact, one of these principles is to show stu-
dents that life obeys the fundamental laws of physics and chemistry. To that end, we
offer chemical explanations of most biochemical reactions, including mechanisms that
tell students how and why things happen.

We are particularly proud of our explanations of oxidation-reduction reactions
since these are extremely important in so many contexts. We describe electron move-
ments in the early chapters, explain reduction potentials in Chapter 10 and use this un-
derstanding to teach about chemiosmotic theory and protonmotive force in Chapter 14
(Electron Transport and ATP Synthesis). The concept is reinforced in the chapter on
photosynthesis.

A focus on biology

While we emphasize chemistry, we also stress the bio in biochemistry. We point out that
biochemical systems evolve and that the reactions that occur in some species are varia-
tions on a larger theme. In this edition, we increase our emphasis on the similarities of

PREFACE XXVii

prokaryotic and eukaryotic systems while we continue to avoid making generalizations
about all organisms based on reactions that occur in a few.

The evolutionary, or comparative, approach to teaching biochemistry focuses at-
tention on fundamental concepts. The evolutionary approach differs in many ways
from other pedagogical methods such as an emphasis on fuel metabolism. The evolu-
tionary approach usually begins with a description of simple fundamental principles or
pathways or processes. These are often the pathways found in bacteria. As the lesson
proceeds, the increasing complexity seen in some other species is explained. At the end
of a chapter we are ready to describe the unique features of the process found in com-
plex multicellular species, such as humans.

Our approach entails additional changes that distinguish us from other textbooks.
When introducing a new chapter, such as lipid metabolism, amino acid metabolism,
and nucleotide metabolism, most other textbooks begin by treating the molecules as
potential food for humans. We start with the biosynthesis pathways since those are the
ones fundamental to all organisms. Then we describe the degradation pathways and end
with an explanation of how they realte to fuel metabolism. This biosynthesis first or-
ganization applies to all the major components of a cell (proteins, nucleotides, nucleic
acids, lipids, amino acids) except carbohydrates where we continue to describe glycoly-
sis ahead of gluconeogenesis. We do, however, emphasize that gluconeogenesis is the
original, primitive pathway and glycolysis evolved later.

This has always been the way DNA replication, transcription, and translation have
been taught. In this book we extend this successful strategy to all the other topics in bio-
chemistry. The chapter on photosynthe sis is an excellent example of how it works in
practice.

In some cases the emphasis on evolution can lead to a profound appreciation of
how complex systems came to exist. Take the citric acid cycle as an example. Students
are often told that such a process cannot be the product of evolution because all the
parts are needed before the cycle can function. We explain in Section 13.9 how such a
pathway can evolve in a stepwise manner.

A focus on accuracy

We are proud of the fact that this is the most scientifically accurate biochemistry text-
book. We have gone to great lengths to ensure that our facts are correct and our explana-
tions of basic concepts reflect the modern consensus among active researchers. Our suc-
cess is due, in large part, to the dedication of our many reviewers and editors.

The emphasis on accuracy means that we check our reactions and our nomencla-
ture against the IUPAC/IUBMB databases. The result is balanced reactions with correct
products and substrates and correct chemical nomenclature. For example, we are one of
the very few textbooks that show all of the citric acid cycle reactions correctly. Previous
editions of this textbook have always scored highly on the Biochemical Howlers website
[bip.cnrs-mrs.fr/bip10/howler.htm] and we feel confident that this edition will achieve a per-
fect score!

We take the time and effort to accurately describe some difficult concepts such as
Gibbs free energy change in a steady-state situation where most reactions are near-
equlibirium reactions (AG = 0). We present correct definitions of the Central Dogma of
Molecular Biology. We don’t avoid genuine areas of scientific controversy such as the
validity of the Three Domain Hypothesis or the mechanism of lysozyme.

A focus on structure-function

Biochemistry is a three-dimensional science. Our inclusion of the latest computer gen-
erated images is intended to clarify the shape and function of molecules and to leave
students with an appreciation for the relationship between the structure and function.
Many of the protein images in this edition are new; they have been skillfully prepared by
Jonathan Parrish of the University of Alberta.

We offer a number of other opportunities. For those students with access to a com-
puter, we have included Protein Data Bank (PDB) reference numbers for the coordinates

from which all protein images were derived. This allows students to further explore the
structures on their own. In addition, we have a gallery of prepared PDB files that stu-
dents can view using Chime or any other molecular viewer; these are posted on the
text’s TheChemistryPlace® website [chemplace.com] as are animations of key dynamic
processes as well as visualization tutorials using Chime.

The emphasis on protein/enzyme structure is a key part of the theme of structure-
function that is one of the most important concepts in biochemistry. At various places
in this new edition we have added material to emphasize this relationship and to develop
it to a greater extent than we have in the past. Some of the most important reactions in
the cell, such as the Q- cycle, cannot be properly understood without understanding the
structure of the enzyme that catalyzes them. Similarly, understanding the properties of
double-stranded DNA is essential to understanding how it serves as the storehouse of
biological information.

Walkthrough of features with some visuals

Interests

Biochemistry is at the root of a number of related sciences, including medicine, forensic
science, biotechnology, and bioengineering; there are many interesting stories to tell.
Throughout the text, you will find boxes that relate biochemistry to other topics. Some
of them are intended to be humorous and help students relate to the material.

BOX 8.1 THE PROBLEM WITH CATS

One of the characteristics of sugars is that they taste sweet.
You certainly know the taste of sucrose and you probably
know that fructose and lactose also taste sweet. So do many
of the other sugars and their derivatives, although we don’t
recommend that you go into a biochemistry lab and start
tasting all the carbohydrates in those white plastic bottles on
the shelves.

Sweetness is not a physical property of molecules. It’s a
subjective interaction between a chemical and taste receptors
in your mouth. There are five different kinds of taste recep-
tors: sweet, sour, salty, bitter, and umami (umami is like the
taste of glutamate in monosodium glutamate). In order to
trigger the sweet taste, a molecule like sucrose has to bind to
the receptor and initiate a response that eventually makes it
to your brain. Sucrose elicits a moderately strong response
that serves as the standard for sweetness. The response to
fructose is almost twice as strong and the response to lactose
is only about one-fifth as strong as that of sucrose. Artificial
sweeteners such as saccharin (Sweet’N Low®), sucralose

(Splenda®), and aspartame (NutraSweet®) bind to the sweet-
ness receptor and cause the sensation of sweetness. They are
hundreds of times more sweet than sucrose.

The sweetness receptor is encoded by two genes called
Tasl r2 and Tasl r3. We don’t know how sucrose and the other
ligands bind to this receptor even though this is a very active
area of research. In the case of sucrose and the artifical sweet-
eners, how can such different molecules elicit the taste of
sweet?

Cats, including lions, tigers and cheetahs, do not have a
functional Taslr2 gene. It has been converted to a pseudo-
gene because of a 247 bp deletion in exon 3. It’s very likely
that your pet cat has never experienced the taste of sweetness.
That explains a lot about cats.

Aspartame

▲ Cats are carnivores. They probably can’t
taste sweetness.

PREFACE XXix

Key Concepts

To help guide students to the information important in each concept, Key Concept
notes have been provided in the margin highlighting this information.

Complete Explanations of the Chemistry

There are thousands of metabolic reactions in a typical organism. You might try to
memorize them all but eventually you will run out of memory. What’s more, memo-
rization will not help you if you encounter something you haven’t seen before. In this
book, we show you some of the basic mechanisms of enzyme -catalyzed reactions — an
extension of what you learned in organic chemistry. If you understand the mechanism,
you’ll understand the chemistry. You’ll have less to memorize, and you’ll retain the in-
formation more effectively.

Margin Notes

There is a great deal of detail in biochemistry but we want you to see both the forest and
the trees. When we need to cross-reference something discussed earlier in the book, or
something that we will come back to later, we put it in the margin. Backward references
offer a review of concepts you may have forgotten. Forward references will help you see
the big picture.

Art

Biochemistry is a three-dimensional science and we have placed a great emphasis on help-
ing you visualize abstract concepts and molecules too small to see. We have tried to make
illustrative figures both informative and beautiful.

KEY CONCEPT

The standard Gibbs free energy change
(A G°') tells us the direction of a reaction
when the concentrations of all products
and reactants are at 1 M concentration.
These conditions will never occur in living
cells. Biochemists are only interested in
actual Gibbs free energy changes (A G),
which are usually close to zero. The
standard Gibbs free energy change (AG°')
tells us the relative concentrations of
reactants and products when the reaction
reaches equilibrium.

The distinction between the normal
flow of information and the Central
Dogma of Molecular Biology is
explained in Section 1.1 and the intro-
duction to Chapter 21.

A-branch

e © V Ferredoxin
^ or
Flavodoxin

P700

Cytochrome c
or

Plastocyanin

XXX PREFACE

Sample Calculations

Sample Calculations are included throughout the text to provide a problem solving
model and illustrate required calculations.

SAMPLE CALCULATION 10.2 Gibbs Free Energy Change

Q: In a rat hepatocyte, the concentrations of ATP, ADP, and the Gibbs free energy change for hydrolysis of ATP in this cell.

Pj are 3.4 mM, 1.3 mM, and 4.8 mM, respectively. Calculate How does this compare to the standard free energy change?

A: The actual Gibbs free energy change is calculated according to Equation 10.10.

[ADP][Pi] [ADP][Pj]

AG rea ction= AG°' reaction + RT In = AG° rea ction + 2.303 RT\og

[ATP] [ATP]

When known values and constants are substituted (with concentrations expressed as molar values), assuming pH7.0 and 25°C.

, , , r (1.3 X 10 _3 )(4.8 X 10 -3 )1

AG = -32000 ] mol” 1 + (8.31 JK“ 1 mol“ 1 )(298 K) 2.303 log

L (3.4 X 10 ) J

AG = -32000 ] mol” 1 + (2480 ] mol -1 ) [2.303 log (1.8 x 10“ 3 )]

AG = -32000 ] mol” 1 - 16000 ] mol” 1

AG = -48000 J mol -1 = -48 kj mol -1

The actual free energy change is about 1 V2 times the standard free energy change.

The Organization

We adopt the metabolism-first strategy of organizing the topics in this book. This means
we begin with proteins and enzymes then describe carbohydrates and lipids. This is fol-
lowed by a description of intermediary metabolism and bioenergetics. The structure of
nucleic acids follows the chapter on nucleotide metabolism and the information flow
chapters are at the back of the book.

While we believe there are significant advantages to teaching the subjects in this
order, we recognize that some instructors prefer to teach information flow earlier in the
course. We have tried to make the last four chapters on nucleic acids, DNA replication,
transcription, and translation less dependant on the earlier chapters but they do discuss
aspects of enzymes that rely on Chapters 4, 5 and 6. Instructors may choose to intro-
duce these last four chapters after a description of enzymes if they wish.

This book has a chapter on coenzymes unlike most other biochemistry textbooks.
We believe that it is important to put more emphasis on the role of coenzymes (and
vitamins) and that’s why we have placed this chapter right after the two chapters on en-
zymes. We know that most instructors prefer to teach the individual coenzymes when
specific examples come up in other contexts. We do that as well. This organization al-
lows instructors to refer back to chapter 7 at whatever point they wish.

Student Supplements

The Study Guide for Principles of Biochemistry

by Scott Lefler

(Arizona State University) and
Allen J. Seism

(Central Missouri State University)

No student should be without this helpful resource. Contents include the following:

• carefully constructed drill problems for each chapter, including short-answer, multiple-
choice, and challenge problems

• comprehensive, step-by-step solutions and explanations for all problems

• a remedial chapter that reviews the general and organic chemistry that students re-
quire for biochemistry — topics are ingeniously presented in the context of a metabolic
pathway

• tables of essential data

PREFACE

XXXI

Chemistry Place for Principles of Biochemistry

An online student tool that includes 3-D modules to help visualize biochemistry and
MediaLabs to investigate important issues related to its particular chapter. Please visit
the site at http://www.chemplace.com.

Acknowledgments

We are grateful to our many talented and thoughtful reviewers who have helped shape this book.

Reviewers who helped in the Fifth Edition:

Accuracy Reviewers

Barry Ganong, Mansfield University

Scott Lefler, Arizona State

Kathleen Nolta, University of Michigan

Content Reviewers

Michelle Chang, University of California, Berkeley

Kathleen Comely, Providence College

Ricky Cox, Murray State University

Michel Goldschmidt- Clermont, University of Geneva

Phil Klebba, University of Oklahoma, Norman

Kristi McQuade, Bradley University

Liz Roberts -Kirchoff, University of Detroit, Mercy

Ashley Spies, University of Illinois

Dylan Taatjes, University of Colorado, Boulder

David Tu, Pennsylvania State University

Jeff Wilkinson, Mississippi State University

Lauren Zapanta, University of Pittsburgh

Reviewers who helped in the Lourth Edition:

Accuracy Reviewers

Neil Haave, University of Alberta
David Watt, University of Kentucky

Content Reviewers

Consuelo Alvarez, Longwood University
Marilee Benore Parsons, University of Michigan
Gary J. Blomquist, University of Nevada, Reno
Albert M. Bobst, University of Cincinnati
Kelly Drew, University of Alaska, Lairbanks
Andrew Leig, Indiana University
Giovanni Gadda, Georgia State University
Donna L. Gosnell, Valdosta State University
Charles Hardin, North Carolina State University
Jane E. Hobson, Kwantlen University College
Ramji L. Khandelwal, University of Saskatchewan
Scott Lefler, Arizona State
Kathleen Nolta, University of Michigan

Jeffrey Schineller, Humboldt State University

Richard Shingles, Johns Hopkins University

Michael A. Sypes, Pennsylvania State University

Martin T. Tuck, Ohio University

Julio F. Turrens, University of South Alabama

David Watt, University of Kentucky

James Zimmerman, Clemson University

Thank you to J. David Rawn who’s work laid the foundation
for this text. We would also like to thank our colleagues who
have previously contributed material for particular chapters
and whose careful work still inhabits this book:

Roy Baker, University of Toronto
Roger W. Brownsey, University of British Columbia
Willy Kalt, Agriculture Canada
Robert K. Murray, University of Toronto
Ray Ochs, St. John’s University
Morgan Ryan, American Scientist
Frances Sharom, University of Guelph
Malcolm Watford, Rutgers, The State University of
New Jersey

Putting this book together was a collaborative effort, and
we would like to thank various members of the team who have
helped give this project life: Jonathan Parrish, Jay McElroy, Lisa
Shoemaker, and the artists of Prentice Hall; Lisa Tarabokjia,
Editorial Assistant, Jessica Neumann, Associate Editor, Lisa
Pierce, Assistant Editor in charge of supplements, Lauren
Layn, Media Editor, Erin Gardner, Marketing Manager; and
Wendy Perez, Production Editor. We would also like to thank
Jeanne Zalesky, our Executive Editor at Prentice Hall.

Finally, we close with an invitation for feedback. Despite
our best efforts (and a terrific track record in the previous edi-
tions), there are bound to be mistakes in a work of this size. We
are committed to making this the best biochemistry text avail-
able; please know that all comments are welcome.

Laurence A. Moran
l.moran@utoronto.ca
Marc D. Perry
marc.perry@utoronto.ca

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About the Authors

Laurence A. Moran

After earning his Ph.D. from Princeton University in 1974,
Professor Moran spent four years at the Universite de Geneve
in Switzerland. He has been a member of the Department of
Biochemistry at the University of Toronto since 1978, special-
izing in molecular biology and molecular evolution. His re-
search findings on heat-shock genes have been published in
many scholarly journals. (l.moran@utoronto.ca)

H. Robert Horton

Dr. Horton, who received his Ph.D. from the University of Mis-
souri in 1962, is William Neal Reynolds Professor Emeritus and
Alumni Distinguished Professor Emeritus in the Department
of Biochemistry at North Carolina State University, where he
served on the faculty for over 30 years. Most of Professor Horton s
research was in protein and enzyme mechanisms.

K. Gray Scrimgeour

Professor Scrimgeour received his doctorate from the Univer-
sity of Washington in 1961 and was a faculty member at the
University of Toronto for over 30 years. He is the author of
The Chemistry and Control of Enzymatic Reactions (1977, Aca-
demic Press), and his work on enzymatic systems has been
published in more than 50 professional journal articles during
the past 40 years. From 1984 to 1992, he was editor of the
journal Biochemistry and Cell Biology. (gray@scrimgeour.ca)

Marc D. Perry

After earning his Ph.D. from the University of Toronto in 1988,
Dr. Perry trained at the University of Colorado, where he stud-
ied sex determination in the nematode C. elegans. In 1994 he
returned to the University of Toronto as a faculty member in
the Department of Molecular and Medical Genetics. His re-
search has focused on developmental genetics, meiosis, and
bioinformatics. In 2008 he joined the Ontario Institute for
Cancer Research. (marc.perry@utoronto.ca)

New problems and solutions for the fifth edition were created by Laurence A. Moran, University of Toronto. The
remaining problems were created by Drs. Robert N. Lindquist, San Francisco State University, Marc Perry,
and Diane M. De Abreu of the University of Toronto.

xxxiii

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Introduction to Biochemistry

B iochemistry is the discipline that uses the principles and language of chemistry
to explain biology. Over the past 100 years biochemists have discovered that the
same chemical compounds and the same central metabolic processes are found
in organisms as distantly related as bacteria, plants, and humans. It is now known that
the basic principles of biochemistry are common to all living organisms. Although sci-
entists usually concentrate their research efforts on particular organisms, their results
can be applied to many other species.

This book is called Principles of Biochemistry because we will focus on the most im-
portant and fundamental concepts of biochemistry — those that are common to most
species. Where appropriate, we will point out features that distinguish particular groups
of organisms.

Many students and researchers are primarily interested in the biochemistry of
humans. The causes of disease and the importance of proper nutrition, for example,
are fascinating topics in biochemistry. We share these interests and that’s why we in-
clude many references to human biochemistry in this textbook. However, we will also
try to interest you in the biochemistry of other species. As it turns out, it is often eas-
ier to understand basic principles of biochemistry by studying many different species
in order to recognize common themes and patterns but a knowledge and appreciation
of other species will do more than help you learn biochemistry. It will also help you
recognize the fundamental nature of life at the molecular level and the ways in which
species are related through evolution from a common ancestor. Perhaps future edi-
tions of this book will include chapters on the biochemistry of life on other planets.
Until then, we will have to be satisfied with learning about the diverse life on our own
planet.

We begin this introductory chapter with a few highlights of the history of biochem-
istry, followed by short descriptions of the chemical groups and molecules you will en-
counter throughout this book. The second half of the chapter is an overview of cell
structure in preparation for your study of biochemistry.

Anything found to be true of E. coli
must also be true of elephants.

— Jacques Monod

Top: Adenovirus. Viruses consist of a nucleic acid molecule surrounded by a protein coat.

2 CHAPTER 1 Introduction to Biochemistry

▲ Friedrich Wohler (1800-1882). Wohler was
one of the founders of biochemistry. By synthe-
sizing urea, Wohler showed that compounds
found in living organisms could be made in
the laboratory from inorganic substances.

▲ Some of the apparatus used by Louis
Pasteur in his Paris laboratory.

▲ Eduard Buchner (1860-1917). Buchner
was awarded the Nobel Prize in Chemistry in
1907 “for his biochemical researches and
his discovery of cell-free fermentation.”

1.1 Biochemistry Is a Modern Science

Biochemistry has emerged as an independent science only within the past 100 years but
the groundwork for the emergence of biochemistry as a modern science was prepared
in earlier centuries. The period before 1900 saw rapid advances in the understanding of
basic chemical principles such as reaction kinetics and the atomic composition of mol-
ecules. Many chemicals produced in living organisms had been identified by the end of
the 19th century. Since then, biochemistry has become an organized discipline and bio-
chemists have elucidated many of the chemical processes of life. The growth of bio-
chemistry and its influence on other disciplines will continue in the 21st century.

In 1828, Friedrich Wohler synthesized the organic compound urea by heating the
inorganic compound ammonium cyanate.

NH 4 (OCN)-^ h 2 n — c — nh 2

This experiment showed for the first time that compounds found exclusively in living or-
ganisms could be synthesized from common inorganic substances. Today we understand
that the synthesis and degradation of biological substances obey the same chemical and
physical laws as those that predominate outside of biology. No special or “vitalistic”
processes are required to explain life at the molecular level. Many scientists date the begin-
nings of biochemistry to Wohlers synthesis of urea, although it would be another 75 years
before the first biochemistry departments were established at universities.

Louis Pasteur (1822-1895) is best known as the founder of microbiology and an
active promoter of germ theory. But Pasteur also made many contributions to biochem-
istry including the discovery of stereoisomers.

Two major breakthroughs in the history of biochemistry are especially notable — the
discovery of the roles of enzymes as catalysts and the role of nucleic acids as informa-
tion-carrying molecules. The very large size of proteins and nucleic acids made their ini-
tial characterization difficult using the techniques available in the early part of the 20th
century. With the development of modern technology we now know a great deal about
how the structures of proteins and nucleic acids are related to their biological functions.

The first breakthrough — identification of enzymes as the catalysts of biological re-
actions — resulted in part from the research of Eduard Buchner. In 1897 Buchner
showed that extracts of yeast cells could catalyze the fermentation of the sugar glucose
to alcohol and carbon dioxide. Previously, scientists believed that only living cells could
catalyze such complex biological reactions.

The nature of biological catalysts was explored by Buchner’s contemporary, Emil
Fischer. Fischer studied the catalytic effect of yeast enzymes on the hydrolysis (break-
down by water) of sucrose (table sugar). He proposed that during catalysis an enzyme
and its reactant, or substrate, combine to form an intermediate compound. He also pro-
posed that only a molecule with a suitable structure can serve as a substrate for a given
enzyme. Fischer described enzymes as rigid templates, or locks, and substrates as
matching keys. Researchers soon realized that almost all the reactions of life are cat-
alyzed by enzymes and a modified lock-and-key theory of enzyme action remains a
central tenet of modern biochemistry.

Another key property of enzyme catalysis is that biological reactions occur much
faster than they would without a catalyst. In addition to speeding up the rates of reac-
tions, enzyme catalysts produce very high yields with few, if any, by-products. In con-
trast, many catalyzed reactions in organic chemistry are considered acceptable with
yields of 50% to 60%. Biochemical reactions must be more efficient because by-
products can be toxic to cells and their formation would waste precious energy. The
mechanisms of catalysis are described in Chapter 5.

The last half of the 20th century saw tremendous advances in the area of structural
biology, especially the structure of proteins. The first protein structures were solved in
the 1950s and 1960s by scientists at Cambridge University (United Kingdom) led by

1.2 The Chemical Elements of Life 3

John C. Kendrew and Max Perutz. Since then, the three-dimensional structures of several
thousand different proteins have been determined and our understanding of the com-
plex biochemistry of proteins has increased enormously. These rapid advances were
made possible by the availability of larger and faster computers and new software that
could carry out the many calculations that used to be done by hand using simple calcu-
lators. Much of modern biochemistry relies on computers.

The second major breakthrough in the history of biochemistry — identification of
nucleic acids as information molecules — came a half-century after Buchner’s and Fis-
cher’s experiments. In 1944 Oswald Avery, Colin MacLeod, and Maclyn McCarty ex-
tracted deoxyribonucleic acid (DNA) from a pathogenic strain of the bacterium
Streptococcus pneumoniae and mixed the DNA with a nonpathogenic strain of the same
organism. The nonpathogenic strain was permanently transformed into a pathogenic
strain. This experiment provided the first conclusive evidence that DNA is the genetic
material. In 1953 James D. Watson and Francis H. C. Crick deduced the three-dimen-
sional structure of DNA. The structure of DNA immediately suggested to Watson and
Crick a method whereby DNA could reproduce itself, or replicate, and thus transmit bi-
ological information to succeeding generations. Subsequent research showed that infor-
mation encoded in DNA can be transcribed to ribonucleic acid (RNA) and then trans-
lated into protein.

The study of genetics at the level of nucleic acid molecules is part of the discipline
of molecular biology and molecular biology is part of the discipline of biochemistry. In
order to understand how nucleic acids store and transmit genetic information, you
must understand the structure of nucleic acids and their role in information flow. You
will find that much of your study of biochemistry is devoted to considering how en-
zymes and nucleic acids are central to the chemistry of life.

As Crick predicted in 1958, the normal flow of information from nucleic acid to
protein is not reversible. He referred to this unidirectional information flow from nu-
cleic acid to protein as the Central Dogma of Molecular Biology. The term “Central
Dogma” is often misunderstood. Strictly speaking, it does not refer to the overall flow of
information shown in the figure. Instead, it refers to the fact that once information in
nucleic acids is transferred to protein it cannot flow backwards from protein to nucleic
acids.

RNA

Translation

Protein

▲ Information flow in molecular biology. The

flow of information is normally from DNA to
RNA. Some RNAs (messenger RNAs) are
translated. Some RNA can be reverse tran-
scribed back to DNA but according Crick’s
Central Dogma of Molecular Biology the
transfer of information from nucleic acid
(e.g., mRNA) to protein is irreversible.

1.2 The Chemical Elements of Life

Six nonmetallic elements — carbon, hydrogen, nitrogen, oxygen, phosphorus, and sul-
fur — account for more than 97% of the weight of most organisms. All these elements
can form stable covalent bonds. The relative amounts of these six elements vary among
organisms. Water is a major component of cells and accounts for the high percentage
(by weight) of oxygen. Carbon is much more abundant in living organisms than in the
rest of the universe. On the other hand, some elements, such as silicon, aluminum, and
iron, are very common in the Earth’s crust but are present only in trace amounts in
cells. In addition to the standard six elements (CHNOPS), there are 23 other elements
commonly found in living organisms (Figure 1.1). These include five ions that are essen-
tial in all species: calcium (Ca©), potassium (K 0 ), sodium (Na 0 ), magnesium (Mg©),
and chloride (Cl®) Note that the additional 23 elements account for only 3% of the
weight of living organisms.

Most of the solid material of cells consists of carbon-containing compounds. The
study of such compounds falls into the domain of organic chemistry. A course in or-
ganic chemistry is helpful in understanding biochemistry because there is considerable
overlap between the two disciplines. Organic chemists are more interested in reactions
that take place in the laboratory, whereas biochemists would like to understand how re-
actions occur in living cells.

Figure 1.2a shows the basic types of organic compounds commonly encountered in
biochemistry. Make sure you are familiar with these terms because we will be using
them repeatedly in the rest of this book.

▲ Emil Fischer (1852-1919). Fischer made
many contributions to our understanding of
the structures and functions of biological
molecules. He received the Nobel Prize in
Chemistry in 1902 “in recognition of the
extraordinary services he has rendered by
his work on sugar and purine synthesis.”

▲ DNA encodes most of the information
required in living cells.

4 CHAPTER 1 Introduction to Biochemistry

IA 0

1.008

1 1 A

IVB

VIB

VI 1 B

\/IIID

IIIA

IVA

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4.003

6.941

9.012

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10.81

12.01

14.01

16.00

19.00

20.18

22.99

24.31

26.98

28.09

30.97

32.07

35.45

39.95

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40.08

44.96

47.87

50.94

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79.90

83.80

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88.91

91.22

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95.94

(98)

101.1

102.9

106.4

107.9

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114.8

118.7

121.8

127.6

126.9

131.3

57*

132.9

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138.9

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180.9

183.8

186.2

190.2

192.2

195.1

197.0

200.6

204.4

207.2

209.0

(209)

(210)

(222)

89**

104

105

106

107

108

109

110

112

113

114

115

116

117

118

(223)

(226)

(227)

(261)

(262)

(263)

(264)

(265)

(268)

(269)

(272)

(277)

(285)

(289)

(293)

58*

140.1

140.9

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(145)

150.4

152.0

157.3

158.9

162.5

164.9

167.3

168.9

173.0

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90**

232.0

231

238.0

(237)

(244)

(243)

(247)

(251)

(252)

100

(257)

101

(258)

102

(259)

103

(262)

▲ Figure 1.1

Periodic Table of the Elements. The important elements found in living cells are shown in color. The
red elements (CHNOPS) are the six abundant elements. The five essential ions are purple. The
trace elements are shown in dark blue (more common) and light blue (less common).

The synthesis of RNA (transcription)
and protein (translation) are described
in Chapters 21 and 22, respectively.

KEY CONCEPT

More than 97% of the weight of most
organisms is made up of only six
elements: carbon, hydrogen, nitrogen,
oxygen, phosphorus, and sulfur
(CHNOPS).

KEY CONCEPT

Living things obey the standard laws of
physics and chemistry. No “vitalistic”
force is required to explain life at the
molecular level.

Biochemical reactions involve specific chemical bonds or parts of molecules called
functional groups (Figure 1.2b). We will encounter several common linkages in bio-
chemistry (Figure 1.2c). Note that all these linkages consist of several different atoms
and individual bonds between atoms. We will learn more about these compounds,
functional groups, and linkages throughout this book. Ester and ether linkages are com-
mon in fatty acids and lipids. Amide linkages are found in proteins. Phosphate ester and
phosphoanhydride linkages occur in nucleotides.

An important theme of biochemistry is that the chemical reactions occurring in-
side cells are the same kinds of reactions that take place in a chemistry laboratory. The
most important difference is that almost all reactions in living cells are catalyzed by en-
zymes and thus proceed at very high rates. One of the main goals of this textbook is to
explain how enzymes speed up reactions without violating the fundamental reaction
mechanisms of organic chemistry.

The catalytic efficiency of enzymes can be observed even when the enzymes and re-
actants are isolated in a test tube. Researchers often find it useful to distinguish between
biochemical reactions that take place in an organism (in vivo) and those that occur
under laboratory conditions (in vitro).

1.3 Many Important Macromolecules Are Polymers

In addition to numerous small molecules, much of biochemistry deals with very large
molecules that we refer to as macromolecules. Biological macromolecules are usually a
form of polymer created by joining many smaller organic molecules, or monomers, via
condensation (removal of the elements of water). In some cases, such as certain carbo-
hydrates, a single monomer is repeated many times; in other cases, such as proteins and
nucleic acids, a variety of different monomers is connected in a particular order. Each
monomer of a given polymer is added by repeating the same enzyme -catalyzed reaction.

1.3 Many Important Macromolecules Are Polymers 5

(a) Organic compounds

R — OH R— C — H

n=o

_73

R— C — OH

◄ Figure 1.2

General formulas of (a) organic compounds,
(b) functional groups, and (c) linkages com-
mon in biochemistry. R represents an alkyl
group (CH 3 (CH 2 ) n ).

Alcohol

Aldehyde

Ketone

Carboxylic acid 1

i 1

R— SH

R — NH 2

R— NH

R — N — R 2

Thiol

(Sulfhydryl)

Primary

Secondary

Tertiary ^

Amines 2

(b) Functional groups

— OH

— C — R

— c —

n=o

Hydroxyl

Acyl

Carbonyl

Carboxylate

— SH

— NH 2 or — NH 3

— O — P — o 0

4-o®

Sulfhydryl

(Thiol)

Amino

0 ®

Phosphate

Phosphoryl

I II I I

— c— o— c— — c— o— c—

I I I

Ester Ether

Amide

1 11 ©

— c— o — p— o u

Phosphate ester

o —

Phosphoanhydride

1 Under most biological conditions,
carboxylic acids exist as carboxylate
anions: O

R— C— O 0

2 Under most biological conditions,
amines exist as ammonium ions:

Ri Ri

R — NH 3 , R — NH 2 and R — NH — R 2

Thus, all of the monomers, or residues, in a macromolecule are aligned in the same di-
rection and the ends of the macromolecule are chemically distinct.

Macromolecules have properties that are very different from those of their con-
stituent monomers. For example, starch is a polymer of the sugar glucose but it is not
soluble in water and does not taste sweet. Observations such as this have led to the gen-
eral principle of the hierarchical organization of life. Each new level of organization re-
sults in properties that cannot be predicted solely from those of the previous level. The
levels of complexity, in increasing order, are: atoms, molecules, macromolecules, or-
ganelles, cells, tissues, organs, and whole organisms. (Note that many species lack one or
more of these levels of complexity. Single-celled organisms, for example, do not have
tissues and organs.) The following sections briefly describe the principal types of
macromolecules and how their sequences of residues or three-dimensional shapes grant
them unique properties.

6 CHAPTER 1 Introduction to Biochemistry

The relative molecular mass ( M r ) of a
molecule is a dimensionless quantity
referring to the mass of a molecule rel-
ative to one-twelfth (1/12) the mass of
an atom of the carbon isotope 12 C.
Molecular weight (M.W.) is another
term for relative molecular mass.

In discussing molecules and macromolecules we will often refer to the molecular
weight of a compound. A more precise term for molecular weight is relative molecular mass
(abbreviated M r ). It is the mass of a molecule relative to one-twelfth (1/12) the mass of an
atom of the carbon isotope 12 C. (The atomic weight of this isotope has been defined as ex-
actly 12 atomic mass units. Note that the atomic weight of carbon shown in the Periodic
Table represents the average of several different isotopes, including 13 C and 14 C.) Because
M r is a relative quantity, it is dimensionless and has no units associated with its value. The
relative molecular mass of a typical protein, for example, is 38,000 (M r = 38,000). The
absolute molecular mass of a compound has the same magnitude as the molecular
weight except that it is expressed in units called daltons (1 dalton = 1 atomic mass unit).
The molecular mass is also called the molar mass because it represents the mass (meas-
ured in grams) of 1 mole, or 6.022 X 10 23 molecules. The molecular mass of a typical
protein is 38,000 daltons, which means that 1 mole weighs 38 kilograms. The main source
of confusion is that the term “molecular weight” has become common jargon in biochem-
istry although it refers to relative molecular mass and not to weight. It is a common error
to give a molecular weight in daltons when it should be dimensionless. In most cases, this
isn’t a very important mistake but you should know the correct terminology.

COO°

H 3 N — C — H

(b) O

H 3 N — CH — C — N — CH — COO u

I I I

R HR

▲ Figure 1 .3

Structure of an amino acid and a dipeptide.

(a) Amino acids contain an amino group
(blue) and a carboxylate group (red). Differ-
ent amino acids contain different side chains
(designated — R). (b) A dipeptide is pro-
duced when the amino group of one amino
acid reacts with the carboxylate group of an-
other to form a peptide bond (red).

KEY CONCEPT

Biochemical molecules are
three-dimensional objects.

A. Proteins

Twenty common amino acids are incorporated into proteins in all cells. Each amino
acid contains an amino group and a carboxylate group, as well as a side chain (R group)
that is unique to each amino acid (Figure 1.3a). The amino group of one amino acid
and the carboxylate group of another are condensed during protein synthesis to form
an amide linkage, as shown in Figure 1.3b. The bond between the carbon atom of one
amino acid residue and the nitrogen atom of the next residue is called a peptide bond.
The end-to-end joining of many amino acids forms a linear polypeptide that may con-
tain hundreds of amino acid residues. A functional protein can be a single polypeptide
or it can consist of several distinct polypeptide chains that are tightly bound to form a
more complex structure.

Many proteins function as enzymes. Others are structural components of cells and
organisms. Finear polypeptides fold into a distinct three-dimensional shape. This shape
is determined largely by the sequence of its amino acid residues. This sequence infor-
mation is encoded in the gene for the protein. The function of a protein depends on its
three-dimensional structure, or conformation.

The structures of many proteins have been determined and several principles gov-
erning the relationship between structure and function have become clear. For example,
many enzymes contain a cleft, or groove, that binds the substrates of a reaction. This
cavity contains the active site of the enzyme — the region where the chemical reaction
takes place. Figure 1.4a shows the structure of the enzyme lysozyme that catalyzes the
hydrolysis of specific carbohydrate polymers. Figure 1.4b shows the structure of the en-
zyme with the substrate bound in the cleft. We will discuss the relationship between
protein structure and function in Chapters 4 and 6.

There are many ways of representing the three-dimensional structures of biopoly-
mers such as proteins. The lysozyme molecule in Figure 1.4 is shown as a cartoon where
the conformation of the polypeptide chain is represented as a combination of wires,
helical ribbons, and broad arrows. Other kinds of representations in the following chap-
ters include images that show the position of every atom. Computer programs that cre-
ate these images are freely available on the Internet and the structural data for proteins
can be retrieved from a number of database sites. With a little practice, any student can
view these molecules on a computer monitor.

B. Polysaccharides

Carbohydrates, or saccharides, are composed primarily of carbon, oxygen, and hydro-
gen. This group of compounds includes simple sugars (monosaccharides) as well as
their polymers (polysaccharides). All monosaccharides and all residues of polysaccha-
rides contain several hydroxyl groups and are therefore polyalcohols. The most com-
mon monosaccharides contain either five or six carbon atoms.

1.3 Many Important Macromolecules Are Polymers 7

Sugar structures can be represented in several ways. For example, ribose (the most
common five-carbon sugar) can be shown as a linear molecule containing four hydroxyl
groups and one aldehyde group (Figure 1.5a). This linear representation is called a Fis-
cher projection (after Emil Fischer). In its usual biochemical form, however, the struc-
ture of ribose is a ring with a covalent bond between the carbon of the aldehyde group
(C-l) and the oxygen of the C-4 hydroxyl group, as shown in Figure 1.5b. The ring form
is most commonly shown as a Haworth projection (Figure 1.5c). This representation is a
more accurate way of depicting the actual structure of ribose. The Haworth projection is
rotated 90° with respect to the Fischer projection and portrays the carbohydrate ring as a
plane with one edge projecting out of the page (represented by the thick lines). However,
the ring is not actually planar. It can adopt numerous conformations in which certain
ring atoms are out-of-plane. In Figure 1.5d, for example, the C-2 atom of ribose lies
above the plane formed by the rest of the ring atoms.

Some conformations are more stable than others so the majority of ribose mole-
cules can be represented by one or two of the many possible conformations. Neverthe-
less, it’s important to note that most biochemical molecules exist as a collection of
structures with different conformations. The change from one conformation to another
does not require the breaking of any covalent bonds. In contrast, the two basic forms of
carbohydrate structures, linear and ring forms, do require the breaking and forming of
covalent bonds.

Glucose is the most abundant six-carbon sugar (Figure 1.6a on page 8). It is the
monomeric unit of cellulose, a structural polysaccharide, and of glycogen and starch,
which are storage polysaccharides. In these polysaccharides, each glucose residue is
joined covalently to the next by a covalent bond between C-l of one glucose molecule
and one of the hydroxyl groups of another. This bond is called a glycosidic bond. In cel-
lulose, C-l of each glucose residue is joined to the C-4 hydroxyl group of the next
residue (Figure 1.6b). The hydroxyl groups on adjacent chains of cellulose interact non-
covalently creating strong, insoluble fibers. Cellulose is probably the most abundant
biopolymer on Earth because it is a major component of flowering plant stems includ-
ing tree trunks. We will discuss carbohydrates further in Chapter 8.

C. Nucleic Acids

Nucleic acids are large macromolecules composed of monomers called nucleotides. The
term polynucleotide is a more accurate description of a single molecule of nucleic acid,
just as polypeptide is a more accurate term than protein for single molecules composed
of amino acid residues. The term nucleic acid refers to the fact that these polynu-
cleotides were first detected as acidic molecules in the nucleus of eukaryotic cells. We

▲ Figure 1.4 Chicken [Gallus gallus) eggwhite
lysozyme, (a) Free lysozyme. Note the char-
acteristic cleft that includes the active site
of the enzyme, (b) Lysozyme with bound
substrate. [PDB 1LZC].

The rules for drawing a molecule as a
Fischer projection are described in
Section 8.1.

Conformations of monosaccharides are
described in more detail in Section 8.3.

Fischer projection Fischer projection

(open-chain form) (ring form)

Haworth projection Envelope conformation

▲ Figure 1.5

Representations of the structure of ribose. (a) In the Fischer projection, ribose is drawn as a linear molecule, (b) In its usual biochemical
form, the ribose molecule is in a ring, shown here as a Fischer projection, (c) In a Haworth projection, the ring is depicted as lying per-
pendicular to the page (as indicated by the thick lines, which represent the bonds closest to the viewer), (d) The ring of ribose is not
actually planar but can adopt 20 possible conformations in which certain ring atoms are out-of-plane. In the conformation shown, C-2 lies
above the plane formed by the rest of the ring atoms.

8 CHAPTER 1 Introduction to Biochemistry

Figure 1.6 ►

Glucose and cellulose, (a) Haworth projection
of glucose, (b) Cellulose, a linear polymer of
glucose residues. Each residue is joined to
the next by a glycosidic bond (red).

The structures of nucleic acids are
described in Chapter 19.

OH H

▲ Figure 1.7

Deoxyribose, the sugar found in deoxyribonu-
cleotides. Deoxyribose lacks a hydroxyl group
at C-2.

The role of ATP in biochemical reac-
tions is described in Section 10.7.

Figure 1.8 ►

Structure of adenosine triphosphate (ATP). The

nitrogenous base adenine (blue) is attached
to ribose (black). Three phosphoryl groups
(red) are also bound to the ribose.

now know that nucleic acids are not confined to the eukaryotic nucleus but are abun-
dant in the cytoplasm and in prokaryotes that don’t have a nucleus.

Nucleotides consist of a five-carbon sugar, a heterocyclic nitrogenous base, and at
least one phosphate group. In ribonucleotides, the sugar is ribose; in deoxyribonu-
cleotides, it is the derivative deoxyribose (Figure 1.7). The nitrogenous bases of nu-
cleotides belong to two families known as purines and pyrimidines. The major purines
are adenine (A) and guanine (G); the major pyrimidines are cytosine (C), thymine (T),
and uracil (U). In a nucleotide, the base is joined to C-l of the sugar, and the phosphate
group is attached to one of the other sugar carbons (usually C-5).

The structure of the nucleotide adenosine triphosphate (ATP) is shown in Figure 1.8.
ATP consists of an adenine moiety linked to ribose by a glycosidic bond. There are three
phosphoryl groups (designated a , /3, and y) esterified to the C-5 hydroxyl group of the ri-
bose. The linkage between ribose and the a-phosphoryl group is a phosphoester linkage
because it includes a carbon and a phosphorus atom, whereas the /3 - and y-phosphoryl
groups in ATP are connected by phosphoanhydride linkages that don’t involve carbon
atoms (see Figure 1.2). All phosphoanhydrides possess considerable chemical potential
energy and ATP is no exception. It is the central carrier of energy in living cells. The potential
energy associated with the hydrolysis of ATP can be used directly in biochemical reactions or
coupled to a reaction in a less obvious way.

In polynucleotides, the phosphate group of one nucleotide is covalently linked to
the C-3 oxygen atom of the sugar of another nucleotide creating a second phosphoester
linkage. The entire linkage between the carbons of adjacent nucleotides is called a phos-
phodiester linkage because it contains two phosphoester linkages (Figure 1.9). Nucleic
acids contain many nucleotide residues and are characterized by a backbone consisting
of alternating sugars and phosphates. In DNA, the bases of two different polynucleotide
strands interact to form a helical structure.

There are several ways of depicting nucleic acid structures depending on which fea-
tures are being described. The ball-and-stick model shown in Figure 1.10 is ideal for show-
ing the individual atoms and the ring structure of the sugars and the bases. In this case, the

OH OH

1.3 Many Important Macromolecules are Polymers 9

two helices can be traced by following the sugar-phosphate backbone emphasized by
the presence of the purple phosphorus atoms surrounded by four red oxygen atoms.
The individual base pairs are viewed edge-on in the interior of the molecule. We will see
several other DNA models in Chapter 19.

RNA contains ribose rather than deoxyribose and it is usually a single-stranded
polynucleotide. There are four different kinds of RNA molecules. Messenger RNA
(mRNA) is involved directly in the transfer of information from DNA to protein. Transfer
RNA (tRNA) is a smaller molecule required for protein synthesis. Ribosomal RNA
(rRNA) is the major component of ribosomes. Cells also contain a heterogeneous class of
small RNAs that carry out a variety of different functions. In Chapters 19 to 22, we will see
how these RNA molecules differ and how their structures reflect their biological roles.

D. Lipids and Membranes

The term “lipid” refers to a diverse class of molecules that are rich in carbon and hydro-
gen but contain relatively few oxygen atoms. Most lipids are not soluble in water but
they do dissolve in some organic solvents. Lipids often have a polar, hydrophilic (water-
loving) head and a nonpolar, hydrophobic (water- fearing) tail (Figure 1.11). In an aque-
ous environment, the hydrophobic tails of such lipids associate while the hydrophobic
heads are exposed to water, producing a sheet called a lipid bilayer. Lipid bilayers form the
structural basis of all biological membranes. Membranes separate cells or compartments
within cells from their environments by acting as barriers that are impermeable to most
water-soluble compounds. Membranes are flexible because lipid bilayers are stabilized by
noncovalent forces.

The simplest lipids are fatty acids — these are long- chain hydrocarbons with a car-
boxylate group at one end. Fatty acids are commonly found as part of larger molecules
called glycerophospholipids consisting of glycerol 3 -phosphate and two fatty acyl groups
(Figure 1.12 on the next page). Glycerophospholipids are major components of biological
membranes.

Other kinds of lipids include steroids and waxes. Steroids are molecules like choles-
terol and many sex hormones. Waxes are common in plants and animals but perhaps
the most familiar examples are beeswax and the wax that forms in your ears.

Membranes are among the largest and most complex cellular structures. Strictly
speaking, membranes are aggregates, not polymers. However, the association of lipid
molecules with each other creates structures that exhibit properties not shown by indi-
vidual component molecules. Their insolubility in water and the flexibility of lipid ag-
gregates give biological membranes many of their characteristics.

◄ Figure 1.9

Structure of a dinucleotide. One deoxyribonu-
cleotide residue contains the pyrimidine
thymine (top), and the other contains the
purine adenine (bottom). The residues are
joined by a phosphodiester linkage between
the two deoxyribose moieties. (The carbon
atoms of deoxyribose are numbered with
primes to distinguish them from the atoms
of the bases thymine and adenine.)

▲ Figure 1.10

Short segment of a DNA molecule. Two differ-
ent polynucleotides associate to form a
double helix. The sequence of base pairs
on the inside of the helix carries genetic
information.

▲ Figure 1 .1 1

Model of a membrane lipid. The molecule
consists of a polar head (blue) and a nonpo-
lar tail (yellow).

Hydrophobic interactions are discussed
in Chapter 2.

10 CHAPTER 1 Introduction to Biochemistry

Figure 1.12 ►

Structures of glycerol 3-phosphate and a glyc-
erophospholipid. (a) The phosphate group of
glycerol 3-phosphate is polar, (b) In a glyc-
erophospholipid, two nonpolar fatty acid
chains are bound to glycerol 3-phosphate
through ester linkages. X represents a sub-
stituent of the phosphate group.

a) 0°

O = P — o e

1 2 3 |

h 2 c — ch — ch 2

HO OH

Glycerol 3-phosphate

Fatty
>- acyl
groups

Glycerophospholipid

(b)

0 = p-

1 2 3 1

h 2 c — ch — ch 2

0 o

1 I

0= c c = o

KEY CONCEPT

Most of the energy required for life is
supplied by light from the sun.

Biological membranes also contain proteins as shown in Figure 1.13. Some of these
membrane proteins serve as channels for the entry of nutrients and the exit of wastes.
Other proteins catalyze reactions that occur specifically at the membrane surface. They
are the sites of many important biochemical reactions. We will discuss lipids and bio-
logical membranes in greater detail in Chapter 9.

1.4 The Energetics of Life

The activities of living organisms do not depend solely on the biomolecules described
in the preceding section and on the multitude of smaller molecules and ions found in
cells. Life also requires the input of energy. Living organisms are constantly transform-
ing energy into useful work to sustain themselves, to grow, and to reproduce. Almost all
this energy is ultimately supplied by the sun.

Lipid

bilayer

▲ Figure 1 .13

General structure of a biological membrane. Biological membranes consist of a lipid bilayer with as-
sociated proteins. The hydrophobic tails of individual lipid molecules associate to form the core of
the membrane. The hydrophilic heads are in contact with the aqueous medium on either side of
the membrane. Most membrane proteins span the lipid bilayer; others are attached to the mem-
brane surface in various ways.

1.4 The Energetics of Life 1 1

Sunlight is captured by plants, algae, and photosynthetic bacteria and used for the
synthesis of biological compounds. Photosynthetic organisms can be ingested as food
and their component molecules used by organisms such as protozoa, fungi, nonphoto-
synthetic bacteria, and animals. These organisms cannot directly convert sunlight into
useful biochemical energy. The breakdown of organic compounds in both photosyn-
thetic and nonphotosynthetic organisms releases energy that can be used to drive the
synthesis of new molecules and macromolecules.

Photosynthesis is one of the key biochemical processes that are essential for life,
even though many species, including animals, benefit only indirectly. One of the by-
products of photosynthesis is oxygen. It is likely that Earth’s atmosphere was trans-
formed by oxygen-producing photosynthetic bacteria during the first several billion
years of its history (a natural example of terraforming). In Chapter 15, we will discuss
the amazing set of reactions that capture sunlight and use it to synthesize biopolymers.

The term metabolism describes the myriad reactions in which organic compounds
are synthesized and degraded and useful energy is extracted, stored, and used. The study
of the changes in energy during metabolic reactions is called bio energetics. Bioenergetics
is part of the field of thermodynamics, a branch of physical science that deals with en-
ergy changes. Biochemists have discovered that the basic thermodynamic principles
that apply to energy flow in nonliving systems also apply to the chemistry of life.

Thermodynamics is a complex and highly sophisticated subject but we don’t need
to master all of its complexities and subtleties in order to understand how it can con-
tribute to an understanding of biochemistry. We will avoid some of the complications
of thermodynamics in this book and concentrate instead on using it to describe some
biochemical principles (discussed in Chapter 10).

A. Reaction Rates and Equilibria

The rate, or speed, of a chemical reaction depends on the concentration of the reac-
tants. Consider a simple chemical reaction where molecule A collides with molecule B
and undergoes a reaction that produces products C and D.

A + B > C + D (1.2)

The rate of this reaction is determined by the concentrations of A and B. At high
concentrations, these reactants are more likely to collide with each other; at low concen-
trations, the reaction might take a long time. We indicate the concentration of a reacting
molecule by enclosing its symbol in square brackets. Thus, [A] means “the concentra-
tion of A” — usually expressed in moles per liter (M). The rate of the reaction is directly
proportional to the product of the concentrations of A and B. This rate can be described
by a proportionality constant, k , that is more commonly called a rate constant.

rate oc [A][B] rate = /c[A][B] (1.3)

Almost all biochemical reactions are reversible. This means that C and D can col-
lide and undergo a chemical reaction to produce A and B. The rate of the reverse reac-
tion will depend on the concentrations of C and D and that rate can be described by a
different rate constant. By convention, the forward rate constant is k\ and the reverse
rate constant is k-\. Reaction 1.4 is a more accurate way of depicting the reaction
shown in Reaction 1.2.

A + B C + D (1.4)

/c_-|

If we begin a test tube reaction by mixing high concentrations of A and B, then the
initial concentrations of C and D will be zero and the reaction will only proceed from
left to right. The rate of the initial reaction will depend on the beginning concentrations
of A and B and the rate constant k\. As the reaction proceeds, the amount of A and B
will decrease and the amount of C and D will increase. The reverse reaction will start to
become significant as the products accumulate. The speed of the reverse reaction will
depend on the concentrations of C and D and the rate constant k-\.

▲ Sunlight on a tropical rain forest. Plants
convert sunlight and inorganic nutrients into
organic compounds.

Inorganic nutrients
(C0 2 , H 2 0)

Light energy

Photosynthetic

organisms

Organic compounds

nergy-^

All organisms

Waste Macromolecules
(C0 2/ H 2 0)

▲ Energy flow. Photosynthetic organisms
capture the energy of sunlight and use it to
synthesize organic compounds. The break-
down of these compounds in both photosyn-
thetic and nonphotosynthetic organisms
generates energy needed for the synthesis of
macromolecules and for other cellular re-
quirements.

12 CHAPTER 1 Introduction to Biochemistry

KEY CONCEPT

The rate of a chemical reaction depends
on the concentrations of the reactants.
The higher the concentration, the faster
the reaction.

At some point, the rates of the forward and reverse reactions will be equal and there
will be no further change in the concentrations of A, B, C, and D. In other words, the re-
action will have reached equilibrium. At equilibrium,

*1 [A] [B] = /C—t [C] [D] (1.5)

KEY CONCEPT

Almost all biochemical reactions are
reversible. When the forward and reverse
reactions are equal, the reaction is at
equilibrium.

In many cases we are interested in the final concentrations of the reactants and
products once the reaction has reached equilibrium. The ratio of product concentra-
tions to reactant concentrations defines the equilibrium constant, K eq . The equilibrium
constant is also equal to the ratio of the forward and reverse rate constants and since Zq
and k_i are constants, so is K eq . Rearranging Equation 1.5 gives,

*1 [C][D]

k - 1 [A][B] eq

( 1 . 6 )

In theory, the concentrations of products and reactants could be identical once the
reaction reaches equilibrium. In that case, K eq = 1 and the forward and reverse rate
constants have the same values. In most cases the value of the equilibrium constant
ranges from 10 -3 to 10 3 meaning that the rate of one of the reactions is much faster
than the other. If K eq = 10 3 then the reaction will proceed mostly to the right and the
final concentrations of C and D will be much higher than the concentrations of A and
B. In this case, the forward rate constant (Zq) will be 1000 times greater than the reverse
rate constant (k-i). This means that collisions between C and D are much less likely to
produce a chemical reaction than collisions between A and B.

▲ Josiah Willard Gibbs (1839-1903). Gibbs
was one of the greatest American scientists
of the 19th century. He founded the modern
field of chemical thermodynamics.

B. Thermodynamics

If we know the energy changes associated with a reaction or process, we can predict the
equilibrium concentrations. We can also predict the direction of a reaction provided we
know the initial concentrations of reactants and products. The thermodynamic quan-
tity that provides this information is the Gibbs free energy (G), named after J. Willard
Gibbs who first described this quantity in 1878.

It turns out that molecules in solution have a certain energy that depends on tem-
perature, pressure, concentration, and other states. The Gibbs free energy change (AG)
for a reaction is the difference between the free energy of the products and the free en-
ergy of the reactants. The overall Gibbs free energy change has two components known
as the enthalpy change (AH, the change in heat content) and the entropy change (AS,
the change in randomness). A biochemical process may generate heat or absorb it from
the surroundings. Similarly, a process may occur with an increase or a decrease in the
degree of disorder, or randomness, of the reactants.

Starting with an initial solution of reactants and products, if the reaction proceeds
to produce more products, then AG must be less than zero (AG < 0). In chemistry
terms, we say that the reaction is spontaneous and energy is released. When AG is
greater than zero (AG > 0), the reaction requires external energy to proceed and it will
not yield more products. In fact, more reactants will accumulate as the reverse reaction
is favored. When AG equals zero (AG = 0), the reaction is at equilibrium; the rates of
the forward and reverse reactions are identical and the concentrations of the products
and reactants no longer change.

We are mostly interested the overall Gibbs free energy change, expressed as

KEY CONCEPT

The Gibbs free energy change (A G) is the
difference between the free energy of the
products of a reaction and that of the
reactants (substrates).

AG = AH - TAS (1.7)

where T is the temperature in Kelvin.

A series of linked processes, such as the reactions of a metabolic pathway in a cell,
usually proceeds only when associated with an overall negative Gibbs free energy
change. Biochemical reactions or processes are more likely to occur, both to a greater
extent and more rapidly, when they are associated with an increase in entropy and a de-
crease in enthalpy.

1.4 The Energetics of Life 13

If we knew the Gibbs free energy of every product and every reactant, it would be a
simple matter to calculate the Gibbs free energy change for a reaction by using Equation 1.8.

Abreaction = AGp roc |ucts — ^^reactants ( 1 - 8 )

Unfortunately, we don’t often know the absolute Gibbs free energies of every bio-
chemical molecule. What we do know are the thermodynamic parameters associated
with the synthesis of these molecules from simple precursors. For example, glucose can
be formed from water and carbon dioxide. We don’t need to know the absolute values of
the Gibbs free energy of water and carbon dioxide in order to calculate the amount of
enthalpy and entropy that are required to bring them together to make glucose. In fact,
the heat released by the reverse reaction (breakdown of glucose to carbon dioxide and
water) can be measured using a calorimeter. This gives us a value for the change in en-
thalpy of synthesis of glucose (AH). The entropy change (A S) for this reaction can also
be determined. We can use these quantities to determine the Gibbs free energy of the re-
action. The true Gibbs free energy of formation AfG is the difference between the ab-
solute free energy of glucose and that of the elements carbon, oxygen and hydrogen.

There are tables giving these Gibbs free energy values for the formation of most bi-
ological molecules. They can be used to calculate the Gibbs free energy change for a re-
action in the same way that we might use absolute values as in Equation 1.9.

AG react j on = AfGp roc | uc t s — AfG reac t an t s (1.9)

In this textbook we will often refer to the AfG value as the Gibbs free energy of a
compound since it can be easily used in calculations as though it were an absolute value.
It can also be called just “Gibbs energy” by dropping the word “free.”

There’s an additional complication that hasn’t been mentioned. For any reaction, in-
cluding the degradation of glucose, the actual free energy change depends on the concen-
trations of reactants and products. Let’s consider the hypothetical reaction in Equation 1.2.
If we begin with a certain amount of A and B and none of the products C and D, then it’s
obvious that the reaction can only go in one direction, at least initially. In thermodynamic
terms, AG react i on is favorable under these conditions. The higher the concentrations of A
and B, the more likely the reaction will occur. This is an important point that we will re-
turn to many times as we learn about biochemistry — the actual Gibbs free energy change
in a reaction depends on the concentrations of the reactants and products.

What we need are some standard values of AG that can be adjusted for concentra-
tion. These standard values are the Gibbs free energy changes measured under certain
conditions. By convention, the standard conditions are 25°C (298 K), 1 atm standard
pressure, and 1.0 M concentration of all products and reactants. In most biochemical
reactions, the concentration of H© is important, and this is indicated by the pH, as will
be described in the next chapter. The standard condition for biochemistry reactions is
pH = 7.0, which corresponds to 10 -7 M H© (rather than 1.0 M as for other reactants
and products). The Gibbs free energy change under these standard conditions is indi-
cated by the symbol AG°'.

The actual Gibbs free energy is related to its standard free energy by

AG a = A C% + R7"ln[A] (1.10)

where R is the universal gas constant (8.315 kj -1 mol -1 ) and T is the temperature in
Kelvin. Gibbs free energy is expressed in units of kj mol -1 . (An older unit is kcal mol -1 ,
which equals 4.184 kj mol -1 .) The term RT ln[A] is sometimes given as 2.303 RT
log [A].

C. Equilibrium Constants and Standard Gibbs Free Energy Changes

For a given reaction, such as that in Reaction 1.2, the actual Gibbs free energy change is
related to the standard free energy change by

[C][D]

AG reac tj on — AG° eac tj on + RT In ^ ^ (1.11)

Thermometer

Insulated

container

Bomb

Water

Sample

▲ The heat given off during a reaction can
be determined by carrying out the reaction
in a sensitive calorimeter.

The importance of the relationship
between A£ and concentration is
explained in Section 10.5.

KEY CONCEPT

The standard Gibbs free energy change
(AG°') tells us the direction of a reaction
when the concentrations of all products
and reactants are at 1 M concentration.
These conditions will never occur in
living cells. Biochemists are only
interested in actual Gibbs free energy
changes (A£), which are usually close to
zero. The standard Gibbs free energy
change (AG°') tells us the relative
concentrations of reactants and products
when the reaction reaches equilibrium.

14 CHAPTER 1 Introduction to Biochemistry

KEY CONCEPT

[C][D]

M=AG°' + R nn—
at equilibrium AG°' + RT In /f eq = 0

KEY CONCEPT

The rate of a reaction is not determined
by the Gibbs free energy change.

If the reaction has reached equilibrium, the ratio of concentrations in the last term of
Equation 1.11 is, by definition, the equilibrium constant (K e q ). When the reaction is at
equilibrium there is no net change in the concentrations of reactants and products, so
the actual Gibbs free energy change is zero ( AG react i on = 0). This allows us to write an
equation relating the standard Gibbs free energy change and the equilibrium constant.
Thus, at equilibrium,

Abreaction = -RT In /C eq = -2.303 RT log K eq (1.12)

This important equation relates thermodynamics and reaction equilibria. Note that
it is the equilibrium constant that is related to the Gibbs free energy change and not the
individual rate constants described in Equations 1.6 and 1.7. It is the ratio of those indi-
vidual rate constants that is important and not their absolute values. The forward and
reverse rates might both be very slow or very fast and still give the same ratio.

D. Gibbs Free Energy and Reaction Rates

Thermodynamic considerations can tell us if a reaction is favored but do not tell how
quickly a reaction will occur. We know, for example, that iron rusts and copper turns
green, but these reactions may take only a few seconds or many years. That’s because,
the rate of a reaction depends on other factors, such as the activation energy.

Activation energies are usually depicted as a hump, or barrier, in diagrams that
show the progress of a reaction from left to right. In Figure 1.14, we plot the Gibbs free
energy at different stages of a reaction as it goes from reactants to products. This
progress is called the reaction coordinate.

The overall change in free energy (AG) can be negative, as shown on the left, or
positive, as shown on the right. In either case, there’s an excess of energy required in
order for the reaction to proceed. The difference between the top of the energy peak and
the energy of the product or reactant with the highest Gibbs free energy is known as the
activation energy ( AG$).

The rate of this reaction depends on the nature of the reaction. Using our example
from Equation 1.2, if every collision between A and B is effective, then the rate is likely
to be fast. On the other hand, if the orientation of individual molecules has to be exactly
right for a reaction to occur then many collisions will be nonproductive and the rate
will be slower. In addition to orientation, the rate depends on the kinetic energy of the
individual molecules. At any given temperature some will be moving slowly when they
collide and they will not have enough energy to react. Others will be moving rapidly
and will carry a lot of kinetic energy.

The activation energy is meant to reflect these parameters. It is a measure of the prob-
ability that a reaction will occur. The activation energy depends on the temperature — it
is lower at higher temperatures. It also depends on the concentration of reactants —
at high concentrations there will be more collisions and the rate of the reaction will be
faster.

The important point is that the rate of a reaction is not predictable from the overall
Gibbs free energy change. Some reactions, such as the oxidation of iron or copper, will
proceed very slowly because their activation energies are high.

Figure 1.14 ►

The progress of a reaction is depicted from left
(reactants) to right (products). In the first dia-
gram, the overall Gibbs free energy change
is negative since the Gibbs free energy of
the products is lower than that of the reac-
tants. In order for the reaction to proceed,
the reactants have to overcome an activation
energy barrier (A Gt). In the second dia-
gram, the overall Gibbs free energy change
for the reaction is positive and the minimum
activation energy is smaller. This means that
the reverse reaction will proceed faster than
the forward reaction.

Reaction coordinate

1.5 Biochemistry and Evolution 15

Most of the reactions that take place inside a cell are very slow in the test tube even
though they are thermodynamically favored. Inside a cell the rates of the normally slow
reactions are accelerated by enzymes. The rates of enzyme -catalyzed reactions can be
10 20 times greater than the rates of the corresponding uncatalyzed reactions. We will
spend some time describing how enzymes work — it is one of the most fascinating top-
ics in biochemistry.

1.5 Biochemistry and Evolution

A famous geneticist, Theodosius Dobzhansky, once said, “Nothing in biology makes
sense except in the light of evolution.” This is also true of biochemistry. Biochemists and
molecular biologists have made major contributions to our understanding of evolution
at the molecular level and the evidence they have uncovered confirms and extends the
data from comparative anatomy, population genetics, and paleontology. We’ve come a
long way from the original evidence of evolution first summarized by Charles Darwin
in the middle of the 19th century.

We now have a very reliable outline of the history of life and the relationships of the
many diverse species in existence today. The first organisms were single cells that we would
probably classify today as prokaryotes. Prokaryotes, or bacteria, do not have a membrane-
bounded nucleus. Fossils of primitive bacteria-like organisms have been found in geologi-
cal formations that are at least 3 billion years old. The modern species of bacteria belong to
such diverse groups as the cyanobacteria, which are capable of photosynthesis, and the
thermophiles, which inhabit hostile environments such as thermal hot springs.

Eukaryotes have cells that possess complex internal architecture, including a promi-
nent nucleus. In general, eukaryotic cells are more complex and much larger than prokary-
otic cells. A typical eukaryotic tissue cell has a diameter of about 25 p, m (25,000 nm),
whereas prokaryotic cells are typically about 1/10 that size. However, evolution has pro-
duced tremendous diversity and extreme deviations from typical sizes are common. For
example, some eukaryotic unicellular organisms are large enough to be visible to the
naked eye and some nerve cells in the spinal columns of vertebrates can be several feet
long. There are also megabacteria that are larger than most eukaryotic cells.

All cells on Earth (prokaryotes and eukaryotes) appear to have evolved from a com-
mon ancestor that existed more than 3 billion years ago. The evidence for common an-
cestry includes the presence in all living organisms of common biochemical building
blocks, the same general patterns of metabolism, and a common genetic code (with
rare, slight variations). We will see many examples of this evidence throughout this
book. The basic plan of the primitive cell has been elaborated on with spectacular in-
ventiveness through billions of years of evolution.

The importance of evolution for a thorough understanding of biochemistry cannot
be overestimated. We will encounter many pathways and processes that only make sense

▲ Charles Darwin (1809-1882). Darwin pub-
lished The Origin of Species in 1859. His
theory of evolution by natural selection ex-
plains adaptive evolution.

◄ Burgess Shale animals. Many transitional
fossils support the basic history of life that
has been worked out over the past few cen-
turies. Pikia, (left) is a primitive chordate
from the time of the Cambrian explosion
about 530 million years ago. These primi-
tive chordates are the ancestors of all mod-
ern chordates, including humans. On the
right is Opabinia, a primitive invertebrate.

16 CHAPTER 1 Introduction to Biochemistry

PROKARYOTES

EUKARYOTES

Gram

Other Proteo- Cyano- positive Cren- Eury-

bacteria bacteria bacteria bacteria archaeota archaeota Animals Fungi Plants Protists

◄ Figure 1.15

The web of life. The two main groups
of prokaryotes are the Eubacteria
(green) and the Archaea (red).
(Adapted from Doolittle (2000).)

when we appreciate that they have evolved from more primitive precursors. The evidence
for evolution at the molecular level is preserved in the sequences of the genes and proteins
that we will study as we learn about biochemistry. In order to fully understand the funda-
mental principles of biochemistry we will need to examine pathways and processes in a
variety of different species including bacteria and a host of eukaryotic model organisms
such as yeast, fruit flies, flowering plants, mice, and humans. The importance of compara-
tive biochemistry has been recognized for over 100 years but its value has increased enor-
mously in the last decade with the publication of complete genome sequences. We are
now able to compare the complete biochemical pathways of many different species.

The relationship of the earliest forms of life can be determined by comparing the
sequences of genes and proteins in modern species. The latest evidence shows that the
early forms of unicellular life exchanged genes frequently giving rise to a complicated
network of genetic relationships. Eventually, the various lineages of bacteria and archae-
bacteria emerged, along with primitive eukaryotes. Further evolution of eukaryotes oc-
curred when they formed a symbiotic union with bacteria, giving rise to mitochondria
and chloroplasts.

The new “web of life” view of evolution (Figure 1.15 ) replaces a more traditional view
that separated prokaryotes into two entirely separate domains called Eubacteria and Ar-
chaea. That distinction is not supported by the data from hundreds of sequenced genomes
so we now see prokaryotes as a single large group with many diverse subgroups, some of
which are shown in the figure. It is also clear that eukaryotes contain many genes that are
more closely related to the old eubacterial groups as well as a minority of genes that are
closer to the old achaeal groups. The early history of life seems to be dominated by rampant
gene exchange between species and this has led to a web of life rather than a tree of life.

Many students are interested in human biochemistry, particularly those aspects of
biochemistry that relate to health and disease. That is an exciting part of biochemistry
but in order to obtain a deep understanding of who we are, we need to know where we
came from. An evolutionary perspective helps explain why we cant make some vitamins

1.7 Prokaryotic Cells: Structural Features

and amino acids and why we have different blood types and different tolerances for
milk products. Evolution also explains the unique physiology of animals, which have
adapted to using other organisms as a source of metabolic fuel.

Every organism is either a single cell or is composed of many cells. Cells exist in a re-
markable variety of sizes and shapes but they can usually be classified as either eukary-
otic or prokaryotic, although some taxonomists continue to split prokaryotes into two
groups: Eubacteria and Archaea.

A simple cell can be pictured as a droplet of water surrounded by a plasma mem-
brane. The water droplet contains dissolved and suspended material including proteins,
polysaccharides, and nucleic acids. The high lipid content of membranes makes them
flexible and self-sealing. Membranes present impermeable barriers to large molecules and
charged species. This property of membranes allows for much higher concentrations of
biomolecules within cells than in the surrounding medium.

The material enclosed by the plasma membrane of a cell is called the cytoplasm.
The cytoplasm may contain large macromolecular structures and subcellular mem-
brane-bound organelles. The aqueous portion of the cytoplasm minus the subcellular
structures is called the cytosol. Eukaryotic cells contain a nucleus and other internal
membrane-bound organelles within the cytoplasm.

Viruses are subcellular infectious particles. They consist of a nucleic acid molecule
surrounded by a protein coat and, in some cases, a membrane. Virus nucleic acid can
contain as few as three genes or as many as several hundred. Despite their biological im-
portance, viruses are not truly cells because they cannot carry out independent meta-
bolic reactions. They propagate by hijacking the reproductive machinery of a host cell
and diverting it to the formation of new viruses. In a sense, viruses are genetic parasites.

There are thousands of different viruses. Those that infect prokaryotic cells are
usually called bacteriophages, or phages. Much of what we know about biochemistry is
derived from the study of viruses and bacteriophages and their interaction with the cells
they infect. For example, introns were first discovered in a human adenovirus like the
one shown on the first page of this chapter and the detailed mapping of genes was first
carried out with bacteriophage T4.

In the following two sections we will explore the structural features of typical
prokaryotic and eukaryotic cells.

Prokaryotes are usually single-celled organisms. The best studied of all living organisms
is the bacterium Escherichia coli (Figure 1.16). This organism has served for half a cen-
tury as a model biological system and many of the biochemical reactions described later
in this book were first discovered in E. coli. E. coli is a fairly typical species of bacteria but
some bacteria are as different from E. coli as we are from diatoms, daffodils and dragonflies.

1.6 The Cell Is the Basic Unit of Life

1.7 Prokaryotic Cells: Structural Features

- Periplasmic space

" Cell wall
Outer membrane

Plasma membrane

◄ Figure 1 .16

Escherichia coli. An E. coli cell is about
0.5 jim in diameter and 1.5 jim long.
Proteinaceous fibers called flagella rotate to
propel the cell. The shorter pili aid in sexual
conjugation and may help E. coli cells
adhere to surfaces. The periplasmic space is
an aqueous compartment separating the
plasma membrane and the outer membrane.

Flagella

18 CHAPTER 1 Introduction to Biochemistry

► Bacteriophage T4. Much of our current un-
derstanding of biochemistry comes from
studies of bacterial viruses such as bacterio-
phage T4.

▲ Max Delbruck and Salvatore Luria. Max Del-
bruck (seated) and Salvatore Luria at the
Cold Spring Harbor Laboratories in 1953.
Delbruck and Luria founded the “phage
group,” a group of scientists who worked on
the genetics and biochemistry of bacteria
and bacteriophage in the 1940s, 1950s,
and 1960s.

Much of this diversity is apparent only at the molecular level. (See Figure 1.15 for the
names of some major groups of prokaryotes.)

Prokaryotes have been found in almost every conceivable environment on Earth,
from hot sulfur springs to beneath the ocean floor to the insides of larger cells. They ac-
count for a significant amount of the biomass on Earth.

Prokaryotes share a number of features in spite of their differences. They lack a nu-
cleus — their DNA is packed in a region of the cytoplasm called the nucleoid region.
Many bacterial species have only 1000 genes. From a biochemists perspective one of the
most fascinating things about bacteria is that, although their chromosomes contain a
relatively small number of genes, they carry out most of the fundamental biochemical
reactions found in all cells, including our own. Hundreds of bacterial genomes have
been completely sequenced and it is now possible to begin to define the minimum
number of enzymes that are consistent with life.

Most bacteria have no internal membrane compartments, although there are many
exceptions. The plasma membrane is usually surrounded by a cell wall made of a rigid
network of covalently linked carbohydrate and peptide chains. This cell wall confers the
characteristic shape of an individual species of bacteria. Despite its mechanical strength,
the cell wall is porous. In addition to the cell wall most bacteria, including E. coli , pos-
sess an outer membrane consisting of lipids, proteins, and lipids linked to polysaccha-
rides. The space between the inner plasma membrane and the outer membrane is called
the periplasmic space. It is the major membrane-bound compartment in bacteria and
plays a crucial role in some important biochemical processes.

Many bacteria have protein fibers, called pili, on their outer surface. The pili serve
as attachment sites for cell-cell interactions. Many species have one or more flagella.
These are long, whip -like structures that can be rotated like the propeller on a boat thus
driving the bacterium through its aqueous environment.

The small size of prokaryotes provides a high ratio of surface area to volume. Sim-
ple diffusion is therefore an adequate means for distributing nutrients throughout the
cytoplasm. One of the prominent macromolecular structures in the cytoplasm is the ri-
bosome — a large RNA-protein complex required for protein synthesis. All living cells
have ribosomes but we will see later that bacterial ribosomes differ from eukaryotic ri-
bosomes in significant details.

1.8 Eukaryotic Cells: Structural Features

Eukaryotes include plants, animals, fungi, and protists. Protists are mostly small, single-
celled organisms that don’t fit into one of the other classes. Along with bacteria these
four groups make up the five kingdoms of life according to one popular classification
scheme. (Older schemes retain the four eukaryotic kingdoms but divide the bacteria
into Eubacteria and Archaea.)

As members of the animal kingdom we are mostly aware of other animals. As rela-
tively large organisms we tend to focus on the large scale. Hence, we know about plants
and mushrooms but not microscopic species.

1.8 Eukaryotic Cells: Structural Features 19

◄ Figure 1.17

The eukaryotic tree of life. The traditional
Plantae, Animalia, and Fungi kingdoms are
branches within the much larger “kingdom”
of Protists.

The latest trees of eukaryotes help us understand the diversity of the protist king-
dom. As shown in Figure 1.17, the animal, plant, and fungal “kingdoms” occupy rela-
tively small branches on the eukaryotic tree of life.

Eukaryotic cells are surrounded by a single plasma membrane unlike bacteria,
which usually have a double membrane. The most obvious feature that distinguishes
eukaryotes from prokaryotes is the presence of a membrane-bound nucleus in eukary-
otes. In fact, eukaryotes are defined by the presence of a nucleus (from the Greek: eu -,
“true” and karuon , “nut” or “kernel .”).

As mentioned earlier, eukaryotic cells are almost always larger than bacterial cells,
commonly 1000-fold greater in volume. Because of their large size complex internal
structures and mechanisms are required for rapid transport and communication both
inside the cell and to and from the external medium. A mesh of protein fibers called the
cytoskeleton extends throughout the cell contributing to cell shape and to the manage-
ment of intracellular traffic.

Almost all eukaryotic cells contain additional internal membrane-bound compart-
ments called organelles. The specific functions of organelles are often closely tied to their
physical properties and structures. Nevertheless, a significant number of specific biochemi-
cal processes occur in the cytosol and the cytosol, like organelles, is highly organized.

The interior of a eukaryotic cell contains an intracellular membrane network. In-
dependent organelles, including the nucleus, mitochondria, and chloroplasts, are em-
bedded in this membrane system that pervades the entire cell. Materials flow within
paths defined by membrane walls and tubules. The intracellular traffic of materials be-
tween compartments is rapid, highly selective, and closely regulated.

Figure 1.18 on the next page shows typical animal and plant cells. Both types have a
nucleus, mitochondria, and a cytoskeleton. Plant cells also contain chloroplasts and vac-
uoles and are often surrounded by a rigid cell wall. Chloroplasts, also found in algae and
some other protists, are the sites of photosynthesis. Plant cell walls are mostly composed
of cellulose, one of the polysaccharides described in Section 1.3B.

Most multicellular eukaryotes contain tissues. Groups of similarly specialized cells
within tissues are surrounded by an extracellular matrix containing proteins and poly-
saccharides. The matrix physically supports the tissue and in some cases directs cell
growth and movement.

KEY CONCEPT

Animals are a relatively small, highly
specialized, branch on the tree of life.

20 CHAPTER 1 Introduction to Biochemistry

(a)

(b)

Endoplasmic
reticulum

Nuclear

envelope

Plasma
membrane

Golgi
apparatus

Vesicles

Chloroplasts

Mitochondrion

Plasma

membrane

Cytoskeleton

Golgi
apparatus

Vesicles

Peroxisome

Nucleus

Vacuole
Cell wall

Peroxisome

Nucleus

Endoplasmic
reticulum

Cytosol

Mitochondrion

Lysosome

▲ Figure 1.18

Eukaryotic cells, (a) Composite animal cell. Animal cells are typical eukaryotic cells containing or-
ganelles and structures also found in protists, fungi, and plants, (b) Composite plant cell. Most
plant cells contain chloroplasts, the sites of photosynthesis in plants and algae; vacuoles, large,
fluid-filled organelles containing solutes and cellular wastes; and rigid cell walls composed mostly
of cellulose.

A. The Nucleus

The nucleus is usually the most obvious structure in a eukaryotic cell. It is structurally de-
fined by the nuclear envelope, a membrane with two layers that join at protein-lined nu-
clear pores. The nuclear envelope is connected to the endoplasmic reticulum (see below).
The nucleus is the control center of the cell containing 95% of its DNA, which is tightly
packed with positively charged proteins called histones and coiled into a dense mass called
chromatin. Replication of DNA and transcription of DNA into RNA occur in the nucleus.
Many eukaryotes have a dense mass in the nucleus called the nucleolus. The nucleolus is a
major site of RNA synthesis and the site of assembly of ribosomes.

Most eukaryotes contain far more DNA than do prokaryotes. Whereas the genetic
material, or genome, of prokaryotes is usually a single circular molecule of DNA, the eu-
karyotic genome is organized as multiple linear chromosomes. In eukaryotes new DNA
and histones are synthesized in preparation for cell division and the chromosomal mate-
rial condenses and separates into two identical sets of chromosomes. This process is
called mitosis (Figure 1.19). The cell is then pinched in two to complete cell division.

Most eukaryotes are diploid — they contain two complete sets of chromosomes.
From time to time eukaryotic cells undergo meiosis resulting in the production of four
haploid cells each with a single set of chromosomes. Two haploid cells — eggs and
sperm, for example — can then fuse to regenerate a typical diploid cell. This process is
one of the key features of sexual reproduction in eukaryotes.

B. The Endoplasmic Reticulum and Golgi Apparatus

A network of membrane sheets and tubules called the endoplasmic reticulum (ER) ex-
tends from the outer membrane of the nucleus. The aqueous region enclosed within the
endoplasmic reticulum is called the lumen. In many cells part of the surface of the
endoplasmic reticulum is coated with ribosomes that are actively synthesizing proteins.

◄ Figure 1.19

Mitosis. The five stages of mitosis are shown. Chromosomes (red) condense and line up in the center
of the cell. Spindle fibers (green) are responsible for separating the recently duplicated chromosomes.

Endoplasmic

reticulum

Cytosol

Ribosomes

Lumen

Nuclear

pore

Nucleus

Nuclear

envelope

As synthesis continues the protein is translocated through the membrane into the
lumen. Proteins destined for export from the cell are completely extruded through the
membrane into the lumen where they are packaged in membranous vesicles. These
vesicles travel through the cell and fuse with the plasma membrane releasing their con-
tents into the extracellular space. The synthesis of proteins destined to remain in the cy-
tosol occurs at ribosomes that are not bound to the endoplasmic reticulum.

A complex of flattened, fluid- filled, membranous sacs called the Golgi apparatus is
often found close to the endoplasmic reticulum and the nucleus. Vesicles that bud off
from the endoplasmic reticulum fuse with the Golgi apparatus. The proteins carried by
the vesicles may be chemically modified as they pass through the layers of the Golgi ap-
paratus. The modified proteins are then sorted, packaged in new vesicles, and trans-
ported to specific destinations inside or outside the cell. The Golgi apparatus was discov-
ered by Camillo Golgi in the 19th century (Nobel Laureate, 1906), although it wasn’t
until many decades later that its role in protein secretion was established.

C. Mitochondria and Chloroplasts

Mitochondria and chloroplasts have central roles in energy transduction. Mitochondria
are the main sites of oxidative energy metabolism. They are found in almost all eukary-
otic cells. Chloroplasts are the sites of photosynthesis in plants and algae.

The mitochondrion has an inner and an outer membrane. The inner membrane is
highly folded, resulting in a surface area three to five times that of the outer membrane.
It is impermeable to ions and most metabolites. The aqueous phase enclosed by the
inner membrane is called the mitochondrial matrix. Many of the enzymes involved in
aerobic energy metabolism are found in the inner membrane and the matrix.

Mitochondria come in many sizes and shapes. The standard jellybean-shaped mi-
tochondrion shown here is found in many cell types but some mitochondria are spher-
ical or have irregular shapes.

The most important role of the mitochondrion is to oxidize organic acids, fatty
acids, and amino acids to carbon dioxide and water. Much of the released energy is con-
served in the form of a proton concentration gradient across the inner mitochondrial
membrane. This stored energy is used to drive the conversion of adenosine diphosphate
(ADP) and inorganic phosphate (Pj) to the energy-rich molecule ATP in a phosphory-
lation process that will be described in detail in Chapter 14. ATP is then used by the cell
for such energy- requiring processes as biosynthesis, transport of certain molecules and
ions against concentration and charge gradients, and generation of mechanical force for
such purposes as locomotion and muscle contraction. The number of mitochondria
found in cells varies widely. Some eukaryotic cells contain only a few mitochondria
whereas others have thousands.

.8 Eukaryotic Cells: Structural Features 21

◄ Nuclear envelope and endoplasmic reticu-
lum (ER) of a eukaryotic cell.

Protein synthesis, sorting, and
secretion are described in Chapter 22.

▲ Golgi apparatus. The Golgi apparatus is re-
sponsible for the modification and sorting of
proteins that have been transported to the
Golgi apparatus by vesicles from the ER.
Vesicles budding off the Golgi apparatus
carry modified material to destinations in-
side and outside the cell.

Outer membrane

membrane

▲ Mitochondrion. Mitochondria are the main
sites of energy transduction in aerobic eu-
karyotic cells. Carbohydrates, fatty acids,
and amino acids are metabolized in this
organelle.

22 CHAPTER 1 Introduction to Biochemistry

► Chloroplast. Chloroplasts are the sites of
photosynthesis in plants and algae. Light
energy is captured by pigments associated
with the thylakoid membrane and used to
convert carbon dioxide and water to carbo-
hydrates.

Outer

Thylakoid

Granum membrane

▲ Micrographs of fluorescently labeled actin
filaments and microtubules in mammalian
cells. (Left) Actin filaments in rat muscle
cells. (Right) Microtubules in human en-
dothelial cells.

Photosynthetic plant cells contain chloroplasts as well as mitochondria. Like mito-
chondria, chloroplasts have an outer membrane and a complex, highly folded, inner
membrane called the thylakoid membrane. Part of the inner membrane forms flattened
sacs called grana (singular, granum). The thylakoid membrane, which is suspended in
the aqueous stroma, contains chlorophyll and other pigments involved in the capture of
light energy. Ribosomes and several circular DNA molecules are also suspended in the
stroma. In chloroplasts the energy captured from light is used to drive the formation of
carbohydrates from carbon dioxide and water.

Mitochondria and chloroplasts are derived from bacteria that entered into internal
symbiotic relationships with primitive eukaryotic cells more than 1 billion years ago.
Evidence for the endosymbiotic ( endo -, “within”) origin of mitochondria and chloro-
plasts includes the presence within these organelles of separate, small genomes and spe-
cific ribosomes that resemble those of bacteria. In recent years scientists have compared
the sequences of mitochondrial and chloroplast genes (and proteins) with those of
many species of bacteria. These studies in molecular evolution have shown that mito-
chondria are derived from primitive members of a particular group of bacteria called
proteobacteria. Chloroplasts are descended from a distantly related class of photosyn-
thetic bacteria called cyanobacteria.

D. Specialized Vesicles

Eukaryotic cells contain specialized digestive vesicles called lysosomes. These vesicles
are surrounded by a single membrane that encloses a highly acidic interior. The acidity
is maintained by proton pumps embedded in the membrane. Lysosomes contain a vari-
ety of enzymes that catalyze the breakdown of cellular macromolecules such as proteins
and nucleic acids. They can also digest large particles such as retired mitochondria and
bacteria ingested by the cell. Lysosomal enzymes are much less active at the near- neutral
pH of the cytosol than they are under the acidic conditions inside the lysosome. The
compartmentalization of lysosomal enzymes keeps them from accidentally catalyzing
the degradation of macromolecules in the cytosol.

Peroxisomes are present in all animal cells and many plant cells. Like lysosomes,
they are surrounded by a single membrane. Peroxisomes carry out oxidation reactions,
some of which produce the toxic compound hydrogen peroxide, (H 2 0 2 ). Some hydro-
gen peroxide is used for the oxidation of other compounds. Excess hydrogen peroxide is
destroyed by the action of the peroxisomal enzyme catalase, which catalyzes the conver-
sion of hydrogen peroxide to water and oxygen.

Vacuoles are fluid-filled vesicles surrounded by a single membrane. They are com-
mon in mature plant cells and some protists. These vesicles are storage sites for water,
ions, and nutrients such as glucose. Some vacuoles contain metabolic waste products
and some contain enzymes that can catalyze the degradation of macromolecules no
longer needed by the plant.

1.9 A Picture of the Living Cell 23

E. The Cytoskeleton

The cytoskeleton is a protein scaffold required for support, internal organization, and even
movement of the cell. Some types of animal cells contain a dense cytoskeleton but it is
much less prominent in most other eukaryotic cells. The cytoskeleton consists of three
types of protein filaments: actin filaments, microtubules, and intermediate filaments. All
three types are built of individual protein molecules that combine to form threadlike fibers.

Actin filaments (also called micro filaments) are the most abundant cytoskeletal
component. They are composed of a protein called actin that forms ropelike threads
with a diameter of about 7 nm. Actin has been found in all eukaryotic cells and is fre-
quently the most abundant protein in the cell. It is also one of the most evolutionarily
conserved proteins. This is evidence that actin filaments were present in the ancestral
eukaryotic cell from which all modern eukaryotes are descended.

Microtubules are strong, rigid fibers frequently packed in bundles. They have a di-
ameter of about 22 nm — much thicker than actin filaments. Microtubules are com-
posed of a protein called tubulin. Microtubules serve as a kind of internal skeleton in
the cytoplasm, but they also form the mitotic spindle during mitosis. In addition, mi-
crotubules can form structures capable of directed movement, such as cilia. The flagella
that propel sperm cells are an example of very long cilia — they are not related to bacter-
ial flagella. The waving motion of cilia is driven by energy from ATP.

Intermediate filaments are found in the cytoplasm of most eukaryotic cells. These
filaments have diameters of approximately 10 nm, which makes them intermediate in
size compared to actin filaments and microtubules. Intermediate filaments line the in-
side of the nuclear envelope and extend outward from the nucleus to the periphery of
the cell. They help the cell resist external mechanical stresses.

1.9 A Picture of the Living Cell

We have now introduced the major structures found within cells and described their
roles. These structures are immense compared to the molecules and polymers that will
be our focus for the rest of this book. Cells contain thousands of different metabolites
and many millions of molecules. In the cytosol of every cell there are hundreds of dif-
ferent enzymes, each acting specifically on only one or possibly a few related metabo-
lites. There may be 100,000 copies of some enzymes per cell but only a few copies of
other enzymes. Each enzyme is bombarded with potential substrates.

Molecular biologist and artist David S. Goodsell has produced captivating images
showing the molecular contents of an E. coli cell magnified 1 million times (Figure 1.20 on
page 26). Approximately 600 cubes of this size represent the volume of the E. coli cell. At
this scale individual atoms are smaller than the dot in the letter i and small metabolites
are barely visible. Proteins are the size of a grain of rice.

A drawing of the molecules in a cell shows how densely packed the cytoplasm can be,
but it cannot give a sense of activity at the atomic scale. All the molecules in a cell are moving
and colliding with each other. The collisions between molecules are fully elastic — the energy
of a collision is conserved in the energy of the rebound. As molecules bounce off each other
they travel a wildly crooked path in space, called the random walk of diffusion. For a small
molecule such as water, the mean distance traveled between collisions is less than the dimen-
sions of the molecule and the path includes many reversals of direction. Despite its convo-
luted path, a water molecule can diffuse the length of an E. coli cell in 1/10 second.

An enzyme and a small molecule will collide 1 million times per second. Under
these conditions, a rate of catalysis typical of many enzymes could be achieved even if
only 1 in about 1000 collisions results in a reaction. Nevertheless, some enzymes cat-
alyze reactions with an efficiency far greater than 1 reaction per 1000 collisions. In fact,
a few enzymes catalyze reactions with almost every molecule of substrate their active
sites encounter — an example of the astounding potency of enzyme- directed chemistry.
The study of the reaction rates of enzymes, or enzyme kinetics, is one of the most fun-
damental aspects of biochemistry. It will be covered in Chapter 6.

Fipids in membranes also diffuse vigorously, though only within the two-dimen-
sional plane of the lipid bilayer. Fipid molecules exchange places with neighboring

▲ Actin. Actin filament showing the organi-
zation in individual subunits of the protein
actin. (Courtesy David S. Goodsell)

MITOCHONDRION

= 200 nm

ANIMAL

CELL

100,000 nm
(100 urn)

RIBOSOME
25 nm

CHLOROPLAST
2000 nm

GLYCOGEN

GRANULE

50 nm

5500 nm

ESCHERICHIA COL I
Flagellum —

15 nm diameter
10,000 nm long

HEMOGLOBIN

= 4 nm

PYRUVATE

DEHYDROGENASE/

25 nm

70S

RIBOSOME

6.0 nm

PLASMA

MEMBRANE

ATP
1.5 nm

WATER MOLECULE
0.4 nm

2.4 nm

AMINO ACID
0.8 nm

6.4 nm

DNA

SUCROSE
1.5 nm

26 CHAPTER 1 Introduction to Biochemistry

▲ Figure 1.20

Portion of the cytosol of an E. coli cell. The

top illustration, in which the contents are
magnified 1 million times, represents a win-
dow 100 x 100 nm. Proteins are in shades
of blue and green. Nucleic acids are in
shades of pink. The large structures are ri-
bosomes. Water and small metabolites are
not shown. The contents in the round inset
are magnified 10 million times, showing
water and other small molecules.

molecules in membranes about 6 million times per second. Some membrane proteins
can also diffuse rapidly within the membrane.

Large molecules diffuse more slowly than small ones. In eukaryotic cells the diffu-
sion of large molecules such as enzymes is retarded even further by the complex net-
work of the cytoskeleton. Large molecules diffuse across a given distance as much as
10 times more slowly in the cytosol than in pure water.

The full extent of cytosolic organization is not yet known. A number of proteins
and enzymes form large complexes that carry out a series of reactions. We will en-
counter several such complexes in our study of metabolism. They are often referred to
as protein machines. This arrangement has the advantage that metabolites pass directly
from one enzyme to the next without diffusing away into the cytosol. Many researchers
are sympathetic to the idea that the cytosol is not merely a random mixture of soluble
molecules but is highly organized in contrast to the long-held impression that simple
solution chemistry governs cytosolic activity. The concept of a highly organized cytosol
is a relatively new idea in biochemistry. It may lead to important new insights about
how cells work at the molecular level.

1.10 Biochemistry Is Multidisciplinary

One of the goals of biochemists is to integrate a large body of knowledge into a molecu-
lar explanation of life. This has been, and continues to be, a challenging task but, in spite
of the challenges, biochemists have made a great deal of progress toward defining and
understanding the basic reactions common to all cells.

The discipline of biochemistry does not exist in a vacuum. We have already seen
how physics, chemistry, cell biology, and evolution contribute to an understanding of
biochemistry. Related disciplines, such as physiology and genetics, are also important.
In fact, many scientists no longer consider themselves to be just biochemists but are also
knowledgeable in several related fields.

Because all aspects of biochemistry are interrelated it is difficult to present one
topic without referring to others. For example, function is intimately related to struc-
ture and the regulation of individual enzyme activities can be appreciated only in the
context of a series of linked reactions. The interrelationship of biochemistry topics is a
problem for both students and teachers in an introductory biochemistry course. The
material must be presented in a logical and sequential manner but there is no universal
sequence of topics that suits every course, or every student. Fortunately, there is general
agreement on the broad outline of an approach to understanding the basic principles of
biochemistry and this textbook follows that outline. We begin with an introductory
chapter on water. We will then describe the structures and functions of proteins and en-
zymes, carbohydrates, and lipids. The third part of the book makes use of structural in-
formation to describe metabolism and its regulation. Finally, we will examine nucleic
acids and the storage and transmission of biological information.

Some courses may cover the material in a slightly different order. For example, the
structures of nucleic acids can be described before the metabolism section. Wherever
possible, we have tried to write chapters so that they can be covered in different orders
in a course depending on the particular needs and interests of the students.

Appendix The Special Terminology of Biochemistry

Most biochemical quantities are specified using Systeme International (SI) units. Some
common SI units are listed in Table 1.1 Many biochemists still use more traditional
units, although these are rapidly disappearing from the scientific literature. For exam-
ple, protein chemists sometimes use the angstrom (A) to report interatomic distances;
1 A is equal to 0.1 nm, the preferred SI unit. Calories (cal) are sometimes used instead of
joules (J); 1 cal is equal to 4.184 J.

The standard SI unit of temperature is the Kelvin, but temperature is most com-
monly reported in degrees Celsius (°C). One degree Celsius is equal in magnitude to
1 Kelvin, but the Celsius scale begins at the freezing point of water (0°C) and 100°C is

Selected Readings 27

TABLE 1.1 SI units commonly used in
biochemistry

Physical

quantity

SI unit

Symbol

Length

meter

Mass

gram

Amount

mole

mol

Volume

liter 0

Energy

joule

Electric potential

volt

Time

second

Temperature

Kelvin* 3

°1 liter = 1 0OO cubic centimeters.
b 273 K = 0° C.

Table 1.2 Prefixes commonly used with
SI units

Prefix

Symbol

Multiplication

factor

giga-

10 9

mega-

10 6

kilo-

10 3

deci-

10- 1

centi-

10“ 2

milli-

10“ 3

micro-

10“ 6

nano-

10“ 9

pico-

icr 12

femto-

io- 15

the boiling point of water at 1 atm. This scale is often referred to as the centigrade scale
( centi - = 1/100). Absolute zero is —273 °C, which is equal to 0 K. In warm-blooded
mammals biochemical reactions occur at body temperature (37°C in humans).

Very large or very small numerical values for some SI units can be indicated by
an appropriate prefix. The commonly used prefixes and their symbols are listed in
Table 1.2. In addition to the standard SI units employed in all fields, biochemistry has
its own special terminology; for example, biochemists use convenient abbreviations for
biochemicals that have long names.

The terms RNA and DNA are good examples. They are shorthand versions of the
long names ribonucleic acid and deoxyribonucleic acid. Abbreviations such as these are
very convenient, and learning to associate them with their corresponding chemical
structures is a necessary step in mastering biochemistry. In this book, we will describe
common abbreviations as each new class of compounds is introduced.

Selected Readings

Chemistry

Bruice, R Y. (2011). Organic Chemistry , 6th ed.
(Upper Saddle River, NJ: Prentice Hall).

Tinoco, I., Sauer, K., Wang, J. C., and Puglisi, J. D.
(2002). Physical Chemistry: Principles and Applica-
tions in Biological Sciences , 4th ed. (Upper Saddle
River, NJ: Prentice Hall).

van Holde, K. E., Johnson, W. C., and Ho, P.S.
(2005). Principles of Physical Biochemistry 2nd ed.
(Upper Saddle River, NJ: Prentice Hall).

Cells

Alberts, B., Bray, D., Hopkin, K., Johnson, A., Lewis,
J., Raff, M., Roberts, K., and Walter, P. (2004).
Essential Cell Biology (New York: Garland).

Lodish, H., Berk, A., Matsudaira, P., Kaiser,

C. A., Kreiger, M., Scott, M. P., Zipursky, L.,
and Darnell, J. (2003). Molecular Cell
Biology , 5th ed. (New York: Scientific
American Books).

Goodsell, D. S. (1993). The Machinery of Life (New
York: Springer- Verlag).

Evolution and the Diversity of Life

Doolittle, W. F. (2000). Uprooting the tree of life.
Sci. Am. 282(2):90-95.

Doolittle, W. F. (2009). Eradicating topological
thinking in prokaryotic systematics and evolution.
Cold Spr. Hbr. Symp. Quant. Biol.

Margulis, L., and Schwartz, K.V. (1998).

Five Kingdoms , 3rd ed. (New York: W.H.

Freeman).

Graur, D., and Li, W.-H. (2000). Fundamentals
of Molecular Evolution (Sunderland, MA:

Sinauer).

Sapp, J. (Ed.) (2005). Microbial Phylogeny and Evo-
lution: Concepts and Controversies. (Oxford, UK:
Oxford University Press).

Sapp, J. (2009) The New Foundations of Evolution.
(Oxford, UK: Oxford University Press).

History of Science

Kohler, R. E. (1975). The History of Biochemistry,
a Survey. /. Hist. Biol 8:275-318.

o o

Water

L ife on Earth is often described as a carbon-based phenomenon but it would be
equally correct to refer to it as a water-based phenomenon. Life probably orig-
inated in water more than three billion years ago and all living cells still de-
pend on water for their existence. Water is the most abundant molecule in most cells
accounting for 60% to 90% of the mass of the cell. The exceptions are cells from which
water is expelled such as those in seeds and spores. Seeds and spores can lie dormant
for long periods of time until they are revived by the reintroduction of water.

Life spread from the oceans to the continents about 500 million years ago. This
major transition in the history of life required special adaptations to enable terrestrial
life to survive in an environment where water was less plentiful. You will encounter
many of these adaptations in the rest of this book.

An understanding of water and its properties is important to the study of biochem-
istry. The macromolecular components of cells — proteins, polysaccharides, nucleic
acids, and lipids — assume their characteristic shapes in response to water. Lor example,
some types of molecules interact extensively with water and, as a result, are very soluble
while other molecules do not dissolve easily in water and tend to associate with each
other in order to avoid water. Much of the metabolic machinery of cells has to operate
in an aqueous environment because water is an essential solvent.

We begin our detailed study of the chemistry of life by examining the properties
of water. The physical properties of water allow it to act as a solvent for ionic and
other polar substances, and the chemical properties of water allow it to form weak
bonds with other compounds, including other water molecules. The chemical proper-
ties of water are also related to the functions of macromolecules, entire cells, and or-
ganisms. These interactions are important sources of structural stability in macro-
molecules and large cellular structures. We will see how water affects the interactions
of substances that have low solubility in water. We will examine the ionization of
water and discuss acid-base chemistry — topics that are the foundation for under-
standing the molecules and processes that we will encounter in subsequent chapters.
It’s important to keep in mind that water is not just an inert solvent; it is also a sub-
strate for many cellular reactions.

There is nothing softer and weaker
than water, And yet there is nothing
better for attacking hard and strong
things. For this reason there is no
substitute for it.

—Lao-Tzu (c. 550 BCE)

▲ Eureka Dunes evening primrose ( Oenothera
californica ) This species only grows in the
sand dunes of Death Valley National Park in
California. It has evolved special mecha-
nisms for conserving water.

Top: Earth from space. The earth is a watery planet and water plays a central role in the chemistry of all life.

2.1 The Water Molecule Is Polar

A water molecule (H 2 0) is V-shaped (Figure 2.1a) and the angle between the two co-
valent (O — H) bonds is 104.5°. Some important properties of water arise from its
angled shape and the intermolecular bonds that it can form. An oxygen atom has
eight electrons and its nucleus has eight protons and eight neutrons. There are two
electrons in the inner shell and six electrons in the outer shell. The outer shell can
potentially accommodate four pairs of electrons in one s orbital and three p orbitals.
However, the structure of water and its properties can be better explained by assum-
ing that the electrons in the outer shell occupy four sp 3 hybrid orbitals. Think of
these four orbitals as occupying the four corners of a tetrahedron that surrounds the
central atom of oxygen. Two of the sp 3 hybrid orbitals contain a pair of electrons and
the other two each contain a single electron. This means that oxygen can form cova-
lent bonds with other atoms by sharing electrons to fill these single electron orbitals.
In water the covalent bonds involve two different hydrogen atoms each of which
shares its single electron with the oxygen atom. In Figure 2.1b each electron is indi-
cated by a blue dot showing that each sp 3 hybrid orbital of the oxygen atom is occu-
pied by two electrons including those shared with the hydrogen atoms. The inner
shell of the hydrogen atom is also filled because of these two shared electrons in the
covalent bond.

The H — O — H bond angle in free water molecules is 104.5° but if the electron or-
bitals were really pointing to the four corners of a tetrahedron, the angle would be
109.5°. The usual explanation for this difference is that there is strong repulsion be-
tween the lone electron pairs and this repulsion pushes the covalent bond orbitals closer
together, reducing the angle from 109.5° to 104.5°.

Oxygen atoms are more electronegative than hydrogen atoms because an oxygen
nucleus attracts electrons more strongly than the single proton in the hydrogen nucleus.
As a result, an uneven distribution of charge occurs within each O — H bond of the
water molecule with oxygen bearing a partial negative charge (8®) and hydrogen bear-
ing a partial positive charge (8®). This uneven distribution of charge within a bond is
known as a dipole and the bond is said to be polar.

The polarity of a molecule depends both on the polarity of its covalent bonds and
its geometry. The angled arrangement of the polar O — H bonds of water creates a per-
manent dipole for the molecule as a whole as shown in Figure 2.2a. A molecule of am-
monia also contains a permanent dipole (Figure 2.2b) Thus, even though water and
gaseous ammonia are electrically neutral, both molecules are polar. The high solubility
of the polar ammonia molecules in water is facilitated by strong interactions with the
polar water molecules. The solubility of ammonia in water demonstrates the principle
that “like dissolves like.”

Not all molecules are polar; for example, carbon dioxide also contains polar cova-
lent bonds but the bonds are aligned with each other and oppositely oriented so the po-
larities cancel each other (Figure 2.2c). As a result, carbon dioxide has no net dipole and
is much less soluble in water than ammonia.

(a)

2 8°

^'o'X

H H

(b)

s©

Bond polarities

35®

(c)

i-r ti

5® H 5

5®

Bond polarities

8° 2 5 ® 8°

0 < =C = 0
Bond polarities

Net dipole

0 = C=0
No net dipole

(a)

O Hydrogen
9 Oxygen

a Figure 2.1 A water molecule, (a) Space-
filling structure of a water molecule.

(b) Angle between the covalent bonds of a
water molecule. Two of the sp 3 hybrid
orbitals of the oxygen atom participate in
covalent bonds with s orbitals of hydrogen
atoms. The other two sp 3 orbitals are
occupied by lone pairs of electrons.

KEY CONCEPT

Polar molecules are molecules with an
unequal distribution of charge so that one
end of the molecules is more negative
and another end is more positive.

◄ Figure 2.2

Polarity of small molecules, (a) The geometry
of the polar covalent bonds of water creates
a permanent dipole for the molecule with
the oxygen bearing a partial negative charge
(symbolized by 28®) and each hydrogen
bearing a partial positive charge (symbolized
by 8®). (b) The pyramidal shape of a mole-
cule of ammonia also creates a permanent
dipole, (c) The polarities of the collinear
bonds in carbon dioxide cancel each other.
Therefore, C0 2 is not polar. (Arrows depict-
ing dipoles point toward the negative charge
with a cross at the positive end.)

30 CHAPTER 2 Water

KEY CONCEPT

Hydrogen bonds form when a hydrogen
atom with a partially positive charge (5®)
is shared between two electronegative
atoms (25®). Hydrogen bonds are much
weaker than covalent bonds.

2.2 Hydrogen Bonding in Water

One of the important consequences of the polarity of the water molecule is that water
molecules attract one another. The attraction between one of the slightly positive hy-
drogen atoms of one water molecule and the slightly negative electron pairs in one of
the sp 3 hybrid orbitals produces a hydrogen bond (Figure 2.3). In a hydrogen bond
between two water molecules the hydrogen atom remains covalently bonded to its oxy-
gen atom, the hydrogen donor. At the same time, it is attracted to another oxygen atom,
called the hydrogen acceptor. In effect, the hydrogen atom is being shared (unequally)
between the two oxygen atoms. The distance from the hydrogen atom to the acceptor
oxygen atom is about twice the length of the covalent bond.

Water is not the only molecule capable of forming hydrogen bonds; these interac-
tions can occur between any electronegative atom and a hydrogen atom attached to an-
other electronegative atom. (We will examine other examples of hydrogen bonding in
Section 2.5B.) Hydrogen bonds are much weaker than typical covalent bonds. The
strength of hydrogen bonds in water and in solutions is difficult to measure directly but
it is estimated to be about 20 kj mol -1 .

H — O — H + H — O — H

O — H

AH f = -20 kJ mol -1 (2J)

About 20 kj mol -1 of heat is given off when hydrogen-bonded water molecules
form in water under standard conditions. (Recall that standard conditions are 1 atm
pressure and a temperature of 25°C.) This value is the standard enthalpy of formation
(AHf). It means that the change in enthalpy when hydrogen bonds form is about -20 kj
per mole of water. This is equivalent to saying that +20 kj mol -1 of heat energy is re-
quired to disrupt hydrogen bonds between water molecules — the reverse of the reaction
shown in Reaction 2.1. This value depends on the type of hydrogen bond. In contrast,
the energy required to break a covalent O — H bond in water is about 460 kj mol -1 , and
the energy required to break a covalent C — H bond is about 410 kj mol -1 . Thus, the
strength of hydrogen bonds is less than 5% of the strength of typical covalent bonds.
Hydrogen bonds are weak interactions compared to covalent bonds.

Orientation is important in hydrogen bonding. A hydrogen bond is most stable when
the hydrogen atom and the two electronegative atoms associated with it (the two oxygen
atoms, in the case of water) are aligned, or nearly in line, as shown in Figure 2.3. Water
molecules are unusual because they can form four O — H — O aligned hydrogen bonds
with up to four other water molecules (Figure 2.4). They can donate each of their two hy-
drogen atoms to two other water molecules and accept two hydrogen atoms from two
other water molecules. Each hydrogen atom can participate in only one hydrogen bond.

The three-dimensional interactions of liquid water are difficult to study but much
has been learned by examining the structure of ice crystals (Figure 2.5). In the common
form of ice, every molecule of water participates in four hydrogen bonds, as expected.
Each of the hydrogen bonds points to the oxygen atom of an adjacent water molecule
and these four adjacent hydrogen-bonded oxygen atoms occupy the vertices of a tetra-
hedron. This arrangement is consistent with the structure of water shown in Figure 2.1

Figure 2.3 ►

Hydrogen bonding between two water mole-
cules. A partially positive (8®) hydrogen
atom of one water molecule attracts the par-
tially negative (25®) oxygen atom of a sec-
ond water molecule, forming a hydrogen
bond. The distances between atoms of two
water molecules in ice are shown. Hydrogen
bonds are indicated by dashed lines high-
lighted in yellow, as shown here and
throughout the book.

0.28 nm

2.2 Hydrogen Bonding in Water

except that the bond angles are all equal (109.5°). This is because the polarity of individual
water molecules, which distorts the bond angles, is canceled by the presence of hydrogen
bonds. The average energy required to break each hydrogen bond in ice has been esti-
mated to be 23 kj mol -1 , making those bonds a bit stronger than those formed in water.

The ability of water molecules in ice to form four hydrogen bonds and the strength
of these hydrogen bonds give ice an unusually high melting point because a large
amount of energy, in the form of heat, is required to disrupt the hydrogen-bonded lat-
tice of ice. When ice melts most of the hydrogen bonds are retained by liquid water.
Each molecule of liquid water can form up to four hydrogen bonds with its neighbors
but most participate in only two or three at any given moment. This means that the
structure of liquid water is less ordered than that of ice. The fluidity of liquid water is
primarily a consequence of the constantly fluctuating pattern of hydrogen bonding as
hydrogen bonds break and re-form. At any given time there will be many water mole-
cules participating in two, three, or four hydrogen bonds with other water molecules.
There will also be many that participate in only one hydrogen bond or none at all. This
is a dynamic structure — the average hydrogen bond lifetime in water is only 10 picosec-
onds (10 -11 s).

The density of most substances increases upon freezing as molecular motion slows
and tightly packed crystals form. The density of water also increases as it cools — until it
reaches a maximum of 1.000 g ml -1 at 4°C (277 K). (This value is not a coincidence.
Grams are defined as the weight of 1 milliliter of water at 4°C.) Water expands as the
temperature drops below 4°C. This expansion is caused by the formation of the more
open hydrogen-bonded ice crystal in which each water molecule is hydrogen-bonded
rigidly to four others. As a result ice is slightly less dense (0.924 g ml -1 ) than liquid
water whose molecules can move enough to pack more closely. Because ice is less dense
than liquid water it floats and water freezes from the top down. This has important bio-
logical implications since a layer of ice on a pond insulates the creatures below from ex-
treme cold.

Two additional properties of water are related to its hydrogen-bonding characteris-
tics — its specific heat and its heat of vaporization. The specific heat of a substance is the
amount of heat needed to raise the temperature of 1 gram of the substance by 1°C. This
property is also called the heat capacity. In the case of water, a relatively large amount of
heat is required to raise the temperature because each water molecule participates in
multiple hydrogen bonds that must be broken in order for the kinetic energy of the
water molecules to increase. The abundance of water in the cells and tissues of all large
multicellular organisms means that temperature fluctuations within cells are minimized.

▲ Figure 2.4

Hydrogen bonding by a water molecule. A

water molecule can form up to four hydro-
gen bonds: the oxygen atom of a water mol-
ecule is the hydrogen acceptor for two hy-
drogen atoms, and each 0 — H group serves
as a hydrogen donor.

▲ Icebergs. Ice floats because it is less
dense than water. However, it is only slightly
less dense than water so most of the mass
of floating ice lies underwater.

◄ Figure 2.5

Structure of ice. Water molecules in ice form
an open hexagonal lattice in which every
water molecule is hydrogen-bonded to four
others. The geometrical regularity of these
hydrogen bonds contributes to the strength
of the ice crystal. The hydrogen-bonding
pattern of ice is more regular than that of
water. The absolute structure of liquid water
has not been determined.

32 CHAPTER 2 Water

BOX 2.1 EXTREME THERMOPHILES

Some species can grow and reproduce at temperatures very
close to 0°C, or even lower. There are cold-blooded fish, for
example, that survive at ocean temperatures below 0°C (salt
lowers the freezing point of water).

At the other extreme are bacteria that live in hot springs
where the average temperature is above 80°C. Some bacteria
inhabit the environment around deep ocean thermal vents
(black smokers) where the average temperature is more than
100°C. (The high pressure at the bottom of the ocean raises
the boiling point of water.)

The record for extreme thermophiles is Strain 121, a
species of archaebacteria that grows and reproduces at
121°C! These extreme thermophiles are among the earliest
branching lineages on the web of life. It’s possible that the
first living cells arose near deep ocean vents.

Deep ocean
hydrothermal
vent. ►

(a) NaCI crystal

O Chlorine

XI X
X if

▲ Figure 2.6

Dissolution of sodium chloride (NaCI) in water.

(a) The ions of crystalline sodium chloride
are held together by electrostatic forces, (b)
Water weakens the interactions between the
positive and negative ions and the crystal
dissolves. Each dissolved Na® and Cl® is
surrounded by a solvation sphere. Only one
layer of solvent molecules is shown. Interac-
tions between ions and water molecules are
indicated by dashed lines.

This feature is of critical biological importance since the rates of most biochemical reac-
tions are sensitive to temperature.

The heat of vaporization of water (-2260 J g -1 ) is also much higher than that of
many other liquids. A large amount of heat is required to convert water from a liquid
to a gas because hydrogen bonds must be broken to permit water molecules to dissoci-
ate from one another and enter the gas phase. Because the evaporation of water
absorbs so much heat, perspiration is an effective mechanism for decreasing body
temperature.

2.3 Water Is an Excellent Solvent

The physical properties of water combine to make it an excellent solvent. We have al-
ready seen that water molecules are polar and this property has important conse-
quences, as we will see below. In addition, water has a low intrinsic viscosity that does
not greatly impede the movement of dissolved molecules. Finally, water molecules
themselves are small compared to some other solvents such as ethanol and benzene.
The small size of water molecules means that many of them can associate with solute
particles to make them more soluble.

A. Ionic and Polar Substances Dissolve in Water

Water can interact with and dissolve other polar compounds and compounds that ion-
ize. Ionization is associated with the gain or loss of an electron, or an H + ion, giving rise
to an atom or a molecule that carries a net charge. Molecules that can dissociate to form
ions are called electrolytes. Substances that readily dissolve in water are said to be
hydrophilic, or water loving. (We will discuss hydrophobic, or water fearing, substances
in the next section.)

Why are electrolytes soluble in water? Recall that water molecules are polar. This
means they can align themselves around electrolytes so that the negative oxygen atoms
of the water molecules are oriented toward the cations (positively charged ions) of the
electrolytes and the positive hydrogen atoms are oriented toward the anions (negatively
charged ions). Consider what happens when a crystal of sodium chloride (NaCI) dis-
solves in water (Figure 2.6) The polar water molecules are attracted to the charged ions
in the crystal. The attractions result in sodium and chloride ions on the surface of the

2.3 Water Is an Excellent Solvent 33

crystal dissociating from one another and the crystal begins to dissolve. Because there
are many polar water molecules surrounding each dissolved sodium and chloride ion,
the interactions between the opposite electric charges of these ions become much weaker
than they are in the intact crystal. As a result of its interactions with water molecules, the
ions of the crystal continue to dissociate until the solution becomes saturated. At this
point, the ions of the dissolved electrolyte are present at high enough concentrations for
them to again attach to the solid electrolyte, or crystallize, and an equilibrium is estab-
lished between dissociation and crystallization.

BOX 2.2 BLOOD PLASMA AND SEAWATER

There was a time when people believed that the ionic compo-
sition of blood plasma resembled that of seawater. This was
supposed to be evidence that primitive organisms lived in the
ocean and land animals evolved a system of retaining the
ocean-like composition of salts.

Careful studies of salt concentrations in the early 20th
century revealed that the concentration of salts in the ocean
were much higher than in blood plasma. Some biochemists
tried to explain this discrepancy by postulating that the com-
position of blood plasma didn’t resemble the seawater of
today but it did resemble the composition of ancient seawa-
ter from several hundred million years ago when multicellu-
lar animals arose.

We now know that the saltiness of the ocean hasn’t
changed very much from the time it first formed over three
billion years ago. There is no direct connection between the
saltiness of blood plasma and seawater. Not only are the overall

v The concentrations of various ions in seawater (blue) and human
blood plasma (red) are compared. Seawater is much saltier and
contains much higher proportions of magnesium and sulfates. Blood
plasma is enriched in bicarbonate (see Section 2.10).

600

500

400

300

200

100

H Seawater
H Blood plasma

■

L □ ,

l H H

Na + K + Mg 2+ Ca + CP SO^f HCO“ 3

concentrations of the major ions (Na + , K + , and CP) very dif-
ferent but the relative concentrations of various other ionic
species are even more different.

The ionic composition of blood plasma is closely mim-
icked by Ringer’s solution, which also contains lactate as a
carbon source. Ringer’s solution can be used as a temporary
substitute for blood plasma when a patient has suffered
blood loss or dehydration.

Blood plasma Ringer's

Na +

140 mM

130 mM

K +

4 mM

103 mM

109 mM

Ca +

2 mM

lactate

5 mM

28 mM

34 CHAPTER 2 Water

▲ Figure 2.7

Structure of glucose. Glucose contains five
hydroxyl groups and a ring oxygen, each of
which can form hydrogen bonds with water.

▲ Figure 2.8

Diffusion, (a) If the cytoplasm were simply
made up of water, a small molecule (red)
would diffuse from one end of a cell to the
other via a random walk, (b) The average time
could be about 10 times longer in a crowded
cytoplasm, with larger molecules (green).

Each dissolved Na® attracts the negative ends of several water molecules whereas
each dissolved Cl® attracts the positive ends of several water molecules (Figure 2.6b).
The shell of water molecules that surrounds each ion is called a solvation sphere and it
usually contains several layers of solvent molecules. A molecule or ion surrounded by
solvent molecules is said to be solvated. When the solvent is water, such molecules or
ions are said to be hydrated.

Electrolytes are not the only hydrophilic substances that are soluble in water. Any
polar molecule will have a tendency to become solvated by water molecules. In addi-
tion, the solubility of many organic molecules is enhanced by formation of hydrogen
bonds with water molecules. Ionic organic compounds such as carboxylates and proto -
nated amines owe their solubility in water to their polar functional groups. Other
groups that confer water solubility include amino, hydroxyl, and carbonyl groups. Mol-
ecules containing such groups disperse among water molecules with their polar groups
forming hydrogen bonds with water.

An increase in the number of polar groups in an organic molecule increases its sol-
ubility in water. The carbohydrate glucose contains five hydroxyl groups and a ring oxy-
gen (Figure 2.7) and is very soluble in water (up to 83 grams of glucose can dissolve in
100 milliliters of water at 17.5°C). Each oxygen atom of glucose can form hydrogen
bonds with water. We will see in other chapters that the attachment of carbohydrates to
some otherwise poorly soluble molecules, including lipids and the bases of nucleosides,
increases their solubility.

B. Cellular Concentrations and Diffusion

The inside of a cell can be very crowded as suggested by David GoodselEs drawings
(Figure 1.17). Consequently, the behavior of solutes in the cytoplasm will be different
from their behavior in a simple solution of water. One of the most important differ-
ences is reduction of the diffusion rate inside cells.

There are three reasons why solutes diffuse more slowly in cytoplasm.

1. The viscosity of cytoplasm is higher than that of water due to the presence of many
solutes such as sugars. This is not an important factor because recent measure-
ments suggest that the viscosity of cytoplasm is only slightly greater than water
even in densely packed organelles.

2. Charged molecules bind transiently to each other inside cells and this restricts their
mobility. These binding effects have a small but significant effect on diffusion rates.

3. Collisions with other molecules inhibit diffusion due to an effect called molecular
crowding. This is the main reason why diffusion is slowed in the cytoplasm.

For small molecules, the diffusion rate inside cells is never more than one-quarter
the rate in pure water. For large molecules, such as proteins, the diffusion rate in the cy-
toplasm may be slowed to about 5% to 10% of the rate in water. This slowdown is due
largely to molecular crowding.

For an individual molecule, the rate of diffusion in water at 20°C is described by
the diffusion coefficient (D 2 o jW ). F° r the protein myoglobin, D 2 o jW = 1 1.3 X 1CT 7 cm 2 s -1 .
From this value we can calculate that the average time to diffuse from one end of a cell
to the other (~10 /mm) is about 0.44 seconds.

But this diffusion time represents the diffusion time in pure water. In the crowed
environment of a typical cell it could take about 10 times longer (4 s). The slower rate is
due to the fact that a protein like myoglobin will be constantly bumping into other large
molecules. Nevertheless, 4 seconds is still a short time. It means that most molecules, in-
cluding smaller metabolites and ions, will encounter each other frequently inside a typ-
ical cell (Figure 2.8). Recent direct measurements of diffusion inside cells reveal that the
effects of molecular crowding are less significant than we used to believe.

C. Osmotic Pressure

If a solvent-permeable membrane separates two solutions that contain different con-
centrations of dissolved substances, or solutes, then molecules of solvent will diffuse
from the less concentrated solution to the more concentrated solution in a process

2.4 Nonpolar Substances Are Insoluble in Water 35

called osmosis. The pressure required to prevent the flow of solvent is called osmotic
pressure. The osmotic pressure of a solution depends on the total molar concentration
of solute, not on its chemical nature.

Water- permeable membranes separate the cytosol from the external medium. The
compositions of intracellular solutions are quite different from those of extracellular
solutions with some compounds being more concentrated and some less concentrated
inside cells. In general, the concentrations of solutes inside the cell are much higher
than their concentrations in the aqueous environment outside the cell. Water molecules
tend to move across the cell membrane in order to enter the cell and dilute the solution
inside the cell. The influx of water causes the cell’s volume to increase but this expan-
sion is limited by the cell membrane. In extreme cases, such as when red blood cells are
diluted in pure water, the internal pressure causes the cells to burst. Some species (e.g.,
plants and bacteria) have rigid cell walls that prevent the membrane expansion. These
cells can develop high internal pressures.

Most cells use several strategies to keep the osmotic pressure from becoming too
great and bursting the cell. One strategy involves condensing many individual mole-
cules into a macromolecule. For example, animal cells that store glucose package it as a
polymer called glycogen which contains about 50,000 glucose residues. If the glucose
molecules were not condensed into a single glycogen molecule the influx of water nec-
essary to dissolve each glucose molecule would cause the cell to swell and burst. Another
strategy is to surround cells with an isotonic solution that negates a net efflux or influx
of water. Blood plasma, for example, contains salts and other molecules that mimic the
osmolarity inside red blood cells (see Box 2.2).

2.4 Nonpolar Substances Are Insoluble in Water

Hydrocarbons and other nonpolar substances have very low solubility in water because
water molecules tend to interact with other water molecules rather than with nonpolar
molecules. As a result, water molecules exclude nonpolar substances forcing them to as-
sociate with each other. For example, tiny oil droplets that are vigorously dispersed in
water tend to coalesce to form a single drop thereby minimizing the area of contact be-
tween the two substances. This is why the oil in a salad dressing separates if you let it sit
for any length of time before putting it on your salad.

Nonpolar molecules are said to be hydrophobic, or water fearing, and this phenome-
non of exclusion of nonpolar substances by water is called the hydrophobic effect. The
hydrophobic effect is critical for the folding of proteins and the self-assembly of biolog-
ical membranes.

The number of polar groups in a molecule affects its solubility in water. Solubility
also depends on the ratio of polar to nonpolar groups in a molecule. For example, one-,
two-, and three-carbon alcohols are miscible with water but larger hydrocarbons
with single hydroxyl groups are much less soluble in water (Table 2.1). In the larger

Table 2.1 Solubilities of short-chain alcohols in water

Alcohol

Structure

Solubility in water
(mol/100 g H 2 0
at 20°C) fl

Methanol

CH 3 OH

Ethanol

ch 3 ch 2 oh

Propanol

CH 3 (CH 2 ) 2 OH

Butanol

CH 3 (CH 2 ) 3 OH

0.11

Pentanol

CH 3 (CH 2 ) 4 OH

0.030

Hexanol

CH 3 (CH 2 ) 5 OH

0.0058

Heptanol

CH 3 (CH 2 ) 6 OH

0.0008

a Infinity (oo) indicates that there is no limit to the solubility of the alcohol in water.

(a) Hypertonic

▲ Hypertonic (a), isotonic (b) and
hypotonic (c) red blood cells.

CHAPTER 2 Water

o=s=o

/ CH2

CH,

/CH2

ch 2

/ CHi

CH 2

/ CH2

CH 2

ch 2

CH,

▲ Figure 2.9

Sodium dodecyl sulfate (SDS), a synthetic
detergent.

molecules, the properties of the nonpolar hydrocarbon portion of the molecule over-
ride those of the polar alcohol group and limit solubility.

Detergents, sometimes called surfactants, are molecules that are both hydrophilic
and hydrophobic. They usually have a hydrophobic chain at least 12 carbon atoms long
and an ionic or polar end. Such molecules are said to be am phi path ic. Soaps, which are
alkali metal salts of long- chain fatty acids are one type of detergent. The soap sodium
palmitate (CH 3 (CH 2 ) 14 COO®Na©), for example, contains a hydrophilic carboxylate
group and a hydrophobic tail. One of the synthetic detergents most commonly used in
biochemistry is sodium dodecyl sulfate (SDS) which contains a 12-carbon tail and a
polar sulfate group (Figure 2.9).

The hydrocarbon portion of a detergent is soluble in nonpolar organic sub-
stances and its polar group is soluble in water. When a detergent is spread on the sur-
face of water a monolayer forms in which the hydrophobic, nonpolar tails of the de-
tergent molecules extend into the air groups of detergent molecules aggregate into
micelles while the hydrophilic, ionic heads are hydrated, extending into the water
(Figure 2.10). When a sufficiently high concentration of detergent is dispersed in
water rather than layered on the surface. In one common form of micelle, the nonpo-
lar tails of the detergent molecules associate with one another in the center of the
structure minimizing contact with water molecules. Because the tails are flexible, the
core of a micelle is liquid hydrocarbon. The ionic heads project into the aqueous solu-
tion and are therefore hydrated. Small, compact micelles may contain about 80 to 100
detergent molecules.

The cleansing action of soaps and other detergents derives from their ability to trap
water- insoluble grease and oils within the hydrophobic interiors of micelles. SDS and
similar synthetic detergents are common active ingredients in laundry detergents. The
suspension of nonpolar compounds in water by their incorporation into micelles is
termed solubilization. Solubilizing nonpolar molecules is a different process than dis-
solving a polar compound. A number of the structures that we will encounter later in
this book, including proteins and biological membranes, resemble micelles in having
hydrophobic interiors and hydrophilic surfaces.

Some dissolved ions such as SCN® (thiocyanate) and C10 4 ® (perchlorate) are
called chaotropes. These ions are poorly solvated compared to ions such as NH4®,
S0 4 2 ®, and H 2 P0 4 ^. Chaotropes enhance the solubility of nonpolar compounds in
water by disordering the water molecules (there is no general agreement on how
chaotropes do this). We will encounter other examples of chaotropic agents such as the
guanidinium ion and the nonionic compound urea when we discuss denaturation and
the three-dimensional structures of proteins and nucleic acids.

▲ Figure 2.10

Cross-sectional views of structures formed by detergents in water. Detergents can form mono-
layers at the air-water interface. They can also form micelles, aggregates of detergent mol-
ecules in which the hydrocarbon tails (yellow) associate in the water-free interior and the
polar head groups (blue) are hydrated.

2.5 Noncovalent Interactions 37

2.5 Noncovalent Interactions < a )

So far in this chapter we have introduced two types of noncovalent interactions —
hydrogen bonds and hydrophobic interactions. Weak interactions such as these play ex-
tremely important roles in determining the structures and functions of macromole-
cules. Weak forces are also involved in the recognition of one macromolecule by
another and in the binding of reactants to enzymes.

There are actually four major noncovalent bonds or forces. In addition to hydrogen
bonds and hydrophobicity there are also charge-charge interactions and van der Waals
forces. Charge-charge interactions, hydrogen bonds, and van der Waals forces are varia-
tions of a more general type of force called electrostatic interactions.

A. Charge-Charge Interactions

Charge-charge interactions are electrostatic interactions between two charged particles.
These interactions are potentially the strongest noncovalent forces and can extend over
greater distances than other noncovalent interactions. The stabilization of NaCl crystals
by interionic attraction between the sodium (Na©) and chloride (Cl©) ions is an ex-
ample of a charge-charge interaction. The strength of such interactions in solution de-
pends on the nature of the solvent. Since water greatly weakens these interactions, the
stability of macromolecules in an aqueous environment is not strongly dependent on ( b )
charge-charge interactions but they do occur. An example of charge-charge interactions
in proteins is when oppositely charged functional groups attract one another. The inter-
action is sometimes called a salt bridge and it’s usually buried deep within the hy-
drophobic interior of a protein where it cant be disrupted by water molecules. The
most accurate term for such interactions is ion pairing.

Charge-charge interactions are also responsible for the mutual repulsion of simi-
larly charged ionic groups. Charge repulsion can influence the structures of individual
biomolecules as well as their interactions with other, like- charged molecules.

In addition to their relatively minor contribution to the stabilization of large mole-
cules, charge-charge interactions play a role in the recognition of one molecule by an-
other. For example, most enzymes have either anionic or cationic sites that bind oppo-
sitely charged reactants.

B. Hydrogen Bonds

Hydrogen bonds, which are also a type of electrostatic interaction, occur in many
macromolecules and are among the strongest noncovalent forces in biological systems.

The strengths of hydrogen bonds such as those between substrates and enzymes and
those between the bases of DNA are estimated to be about 25-30 kj mol -1 . These hydro-
gen bonds are a bit stronger than those formed between water molecules (Section 2.2).
Hydrogen bonds in biochemical molecules are strong enough to confer structural sta-
bility but weak enough to be broken readily.

In general, when a hydrogen atom is covalently bonded to a strongly elec-
tronegative atom, such as nitrogen, oxygen, or sulfur, a hydrogen bond can only
form when the hydrogen atom lies approximately 0.2 nm from another strongly
electronegative atom with an unshared electron pair. As previously described in
the case of hydrogen bonds between water molecules the covalently bonded atom
(designated D in Figure 2.11a) is the hydrogen donor and the atom that attracts the
proton (designated A in Figure 2.1 la) is the hydrogen acceptor. The total distance be-
tween the two electronegative atoms participating in a hydrogen bond is typically be-
tween 0.27 nm and 0.30 nm. Some common examples of hydrogen bonds are shown
in Figure 2.11b.

A hydrogen bond has many of the characteristics of a covalent bond but it is much
weaker. You can think of a hydrogen bond as a partial sharing of electrons. (Recall that
in a true covalent bond a pair of electrons is shared between two atoms.) The three atoms
involved in a hydrogen bond are usually aligned to form a straight line where the center
of the hydrogen atoms falls directly on a line drawn between the two electronegative

▲ Salt bridges, (a) One kind of salt bridge,
(b) Another kind of salt bridge.

38 CHAPTER 2 Water

(a)

<EHa) ®=

Covalent Hydrogen
bond bond

-0.1 nm -0.2 nm

(b)

,0 — H

-0=C V

N // \ /

c — c c — c

// \ // \

-c N — H-—N c — H

\ / \ /

N=C C— N

\ // \

N — H O R

Guanine H

Cytosine

Figure 2.12 ▲

Hydrogen bonding between the complementary bases guanine and cytosine in DNA.

/O-H — \

\ /

N _ Ha ____ l0 =c

/ \

\ /

IN, — H a — 1 0

/ \

\ S

IN, — H ■ — ■ N

/ \

▲ Figure 2.1 1

Hydrogen bonds, (a) Hydrogen bonding be-
tween a — D — H group (the hydrogen donor)
and an electronegative atom A — (the hydro-
gen acceptor). A typical hydrogen bond is ap-
proximately 0.2 nm long, roughly twice the
length of the covalent bond between hydrogen
and nitrogen, oxygen, or sulfur. The total dis-
tance between the two electronegative atoms
participating in a hydrogen bond is therefore
approximately 0.3 nm. (b) Examples of bio-
logically important hydrogen bonds.

Hydrogen bonding between base pairs
in double-stranded DNA makes only a
small contribution to the stability of
DNA, as described in Section 19.2C.

KEY CONCEPT

Hydrogen bonds between and within
biological molecules are easily disrupted
by competition with water molecules.

atoms. Small deviations from this alignment are permitted but such hydrogen bonds are
weaker than the standard form.

All of the functional groups shown in Figure 2.11 are also capable of forming hy-
drogen bonds with water molecules. In fact, when they are exposed to water they are far
more likely to interact with water molecules because the concentration of water is so
high. In order for hydrogen bonds to form between, or within, biochemical macromol-
ecules the donor and acceptor groups have to be shielded from water. In most cases, this
shielding occurs because the groups are buried in the hydrophobic interior of the
macromolecule where water cant penetrate. In DNA, for example, the hydrogen bonds
between complementary base pairs are in the middle of the double helix (Figure 2.12).

C. Van der Waals Forces

The third weak force involves the interactions between permanent or transient dipoles
of two molecules. These forces are of short range and small magnitude, about 13 kj
mol -1 and 0.8 kj mol -1 , respectively.

These electrostatic interactions are called van der Waals forces named after the
Dutch physicist Johannes Diderik van der Waals. They only occur when atoms are very
close together. Van der Waals forces involve both attraction and repulsion. The attrac-
tive forces, also known as London dispersion forces, originate from the infinitesimal di-
pole generated in atoms by the random movement of the negatively charged electrons
around the positively charged nucleus. Thus, van der Waals forces are dipolar, or elec-
trostatic, attractions between the nuclei of atoms or molecules and the electrons of
other atoms or molecules. The strength of the interaction between the transiently in-
duced dipoles of nonpolar molecules such as methane is about 0.4 kj mol -1 at an inter-
nuclear separation of 0.3 nm. Although they operate over similar distances, van der
Waals forces are much weaker than hydrogen bonds.

There is also a repulsive component to van der Waals forces. When two atoms are
squeezed together the electrons in their orbitals repel each other. The repulsion in-
creases exponentially as the atoms are pressed together and at very close distances it be-
comes prohibitive.

The sum of the attractive and repulsive components of van der Waals forces yields
an energy profile like that in Figure 2.13. At large intermolecular distances the two atoms
do not interact and there are no attractive or repulsive forces between them. As the atoms
approach each other (moving toward the left in the diagram) the attractive force in-
creases. This attractive force is due to the delocalization of the electron cloud around the
atoms. You can picture this as a shift in electrons around one of the atoms such that the
electrons tend to localize on the side opposite that of the other approaching atom. This
shift creates a local dipole where one side of the atom has a slight positive charge and the
other side has a slight negative charge. The side with the small positive charge attracts the
other negatively charged atom. As the atoms move even closer together the effect of this
dipole diminishes and the overall influence of the negatively charged electron cloud be-
comes more important. At short distances the atoms repel each other.

2.6 Water is Nucleophilic 39

The optimal packing distance is the point at which the attractive forces are maxi-
mized. This distance corresponds to the energy trough in Figure 2.13 and it is equal to
the sum of the van der Waals radii of the two atoms. When the atoms are separated by
the sum of their two van der Waals radii they are said to be in van der Waals contact, p
Typical van der Waals radii of several atoms are shown in Table 2.2. £

In some cases, the shift in electrons is influenced by the approach of another atom. m
This is an induced dipole. In other cases, the delocalization of electrons is a permanent
feature of the molecule as we saw in the case of water (Section 2.1). These permanent
dipoles also give rise to van der Waals forces.

Although individual van der Waals forces are weak, the clustering of atoms
within a protein, nucleic acid, or biological membrane permits formation of a large
number of these weak interactions. Once formed, these cumulative weak forces play
important roles in maintaining the structures of the molecules. For example, the het-
erocyclic bases of nucleic acids are stacked one above another in double-stranded
DNA. This arrangement is stabilized by a variety of noncovalent interactions, espe-
cially van der Waals forces. These forces are collectively known as stacking interac-
tions (see Chapter 19).

D. Hydrophobic Interactions

The association of a relatively nonpolar molecule or group with other nonpolar molecules
is termed a hydrophobic interaction. Although hydrophobic interactions are sometimes
called hydrophobic “bonds? this description is incorrect. Nonpolar molecules don’t aggre-
gate because of mutual attraction but because the polar water molecules surrounding them
tend to associate with each other rather than with the nonpolar molecules (Section 2.4).
For example, micelles (Figure 2.10) are stabilized by hydrophobic interactions.

The hydrogen-bonding pattern of water is disrupted by the presence of a nonpolar
molecule. Thus, water molecules surrounding a less polar molecule in solution are more
restricted in their interactions with other water molecules. These restricted water mole-
cules are relatively immobile, or ordered, in the same way that molecules at the surface
of water are ordered in the familiar phenomenon of surface tension. However, water
molecules in the bulk solvent phase are much more mobile, or disordered. In thermo-
dynamic terms, there is a net gain in the combined entropy of the solvent and the non-
polar solute when the nonpolar groups aggregate and water is freed from its ordered
state surrounding the nonpolar groups.

Hydrophobic interactions, like hydrogen bonds, are much weaker than covalent bonds
but stronger than van der Waals interactions. For example, the energy required to transfer a
— CH 2 — group from a hydrophobic to an aqueous environment is about 3 kj mol -1 .

Although individual hydrophobic interactions are weak, the cumulative effect of
many hydrophobic interactions can have a significant effect on the stability of a macro -
molecule. The three-dimensional structure of most proteins, for example, is largely de-
termined by hydrophobic interactions formed during the spontaneous folding of the
polypeptide chain. Water molecules are bound to the outside surface of the protein but
can’t penetrate the interior where most of the nonpolar groups are located.

All four of the interactions covered here are individually weak compared to cova-
lent bonds but the combined effect of many such weak interactions can be quite
strong. The most important noncovalent interactions in biomolecules are shown in
Figure 2.14.

▲ Figure 2.13

Effect of internuclear separation on van der
Waals forces. Van der Waals forces are
strongly repulsive at short internuclear dis-
tances and very weak at long internuclear
distances. When two atoms are separated by
the sum of their van der Waals radii, the van
der Waals attraction is maximal.

Table 2.2 Van der Waals radii of several
atoms

Atom

Radius (nm)

Hydrogen

0.12

Oxygen

0.14

Nitrogen

0.15

Carbon

0.17

Sulfur

0.18

Phosphorus

0.19

KEY CONCEPT

Weak interactions are individually weak
but the combined effect of a large number
of weak interactions is a significant
organizing force.

2.6 Water Is Nucleophilic

In addition to its physical properties, the chemical properties of water are also impor-
tant in biochemistry because water molecules can react with biological molecules. The
electron- rich oxygen atom determines much of water’s reactivity in chemical reactions.
Electron-rich chemicals are called nucleophiles (nucleus lovers) because they seek posi-
tively charged (electron-deficient) species called electrophiles (electron lovers). Nucle-
ophiles are either negatively charged or have unshared pairs of electrons. They attack

40 CHAPTER 2 Water

/; ©

— C;G H 3 N —

Charge-charge interaction
~40 to 200 kJ moH

\ /

C=0 H— N

/ \

Hydrogen bond
~25 to 30 kJ mol -1

H H

I I

— C— H H — C —

I I

H H

I I

— C— H H — C —

van der Waals interaction
~0.4 to 4kJ mol -1

\ /

ch 2 h 2 c

Hydrophobic interaction
-3 to 1 0 kJ mol -1

▲ Figure 2.14

Typical noncovalent interactions in biomole-
cules. Charge-charge interactions, hydrogen
bonds, and van der Waals interactions are
electrostatic interactions. Hydrophobic inter-
actions depend on the increased entropy of
the surrounding water molecules rather than
on direct attraction between nonpolar
groups. For comparison, the dissociation en-
ergy for a covalent bond such as C — H or
C — C is approximately 340-450 kJ mol -1 .

R O

CO 1 11

^h 3 n — ch— c — nh— ch — c

+ H,0

Condensation

Hydrolysis

I /° /°

@ H 3 N — CH — C X + @ H 3 N — CH — C 7

Figure 2.15 ▲

Hydrolysis of a peptide. In the presence of water the peptide bonds in proteins and peptides are
hydrolyzed. Condensation, the reverse of hydrolysis, is not thermodynamically favored.

electrophiles during substitution or addition reactions. The most common nucleophilic
atoms in biology are oxygen, nitrogen, sulfur, and carbon.

The oxygen atom of water has two unshared pairs of electrons making it nucle-
ophilic. Water is a relatively weak nucleophile but its cellular concentration is so high
that one might reasonably expect it to be very reactive. Many macromolecules should be
easily degraded by nucleophilic attack by water. This is, in fact, a correct expectation.
Proteins, for example, are hydrolyzed, or degraded, by water to release their monomeric
units, amino acids (Figure 2.15). The equilibrium for complete hydrolysis of a protein
lies far in the direction of degradation; in other words, the ultimate fate of all proteins is
destruction by hydrolysis!

If there is so much water in cells then why aren’t all biopolymers rapidly degraded?
Similarly, if the equilibrium lies toward breakdown, how does biosynthesis occur in an
aqueous environment? Cells avoid these problems in several ways. For example, the
linkages between the monomeric units of macromolecules, such as the peptide bonds in
proteins and the ester linkages in DNA, are relatively stable in solution at cellular pH
and temperature in spite of the presence of water. In this case, the stability of linkages
refers to their rate of hydrolysis in water and not their thermodynamic stability.

The chemical properties of water combined with its high concentration mean that
the Gibbs free energy change for hydrolysis (AG) is negative. This means that all hydrol-
ysis reactions are thermodynamically favorable. However, the rate of the reactions in-
side the cell is so slow that macromolecules are not appreciably degraded by sponta-
neous hydrolysis during the average lifetime of a cell. It is important to keep in mind the
distinction between the preferred direction of a reaction, as indicated by the Gibbs free
energy change, and the rate of the reaction, as indicated by the rate constant (Section
1.4D). The key concept is that because of the activation energy there is no direct corre-
lation between the rate of a reaction and the final equilibrium values of the reactants
and products.

Cells can synthesize macromolecules in an aqueous environment even though
condensation reactions — the reverse of hydrolysis — are thermodynamically unfavor-
able. They do this by using the chemical potential energy of ATP to overcome an unfa-
vorable thermodynamic barrier. Furthermore, the enzymes that catalyze such reactions
exclude water from the active site where the synthesis reactions occur. These reactions
usually follow two-step chemical pathways that differ from the reversal of hydrolysis.
For example, the simple condensation pathway shown in Figure 2.15 is not the path-
way that is used in living cells because the presence of high concentrations of water
makes the direct condensation reaction extremely unfavorable. In the first synthetic
step, which is thermodynamically uphill, the molecule to be transferred reacts with
ATP to form a reactive intermediate. In the second step, the activated group is readily

2.7 Ionization of Water 41

BOX 2.3 THE CONCENTRATION
OF WATER

The density of water varies with tempera-
ture. It is defined as 1.00000 g/ml at
3.98°C. The density is 0.99987 at 0°C and
0.99707 at 25°C.

The molecular mass of the most
common form of water is M r =18.01056.
The concentration of pure water at
3.98°C is 55.5 M (1000 | 18.01).

Many biochemical reactions in-
volve water as either a reactant or a
product and the high concentration of
water will affect the equilibrium of the
reaction.

KEY CONCEPT

There is a difference between the rate of
a reaction and whether it is
thermodynamically favorable. Biological
molecules are stable because the rate of
spontaneous hydrolysis is slow.

transferred to the attacking nucleophile. In Chapter 22 we will see that the reactive in-
termediate in protein synthesis is an amino acyl -tRNA that is formed in a reaction in-
volving ATP. The net result of the biosynthesis reaction is to couple the condensation
to the hydrolysis of ATP.

The role of ATP in coupled reactions is
described in Section 10.7.

2.7 Ionization of Water

One of the important properties of water is its slight tendency to ionize. Pure water
contains a low concentration of hydronium ions (H 3 0®) and an equal concentration of
hydroxide ions (OH®). The hydronium and hydroxide ions are formed by a nucleophilic
attack of oxygen on one of the protons in an adjacent water molecule.

o— H <-

H,0 + H,0

1 ©

. O . + ^o — H

h^©^h

H 3 0® + OH°

( 2 . 2 )

The red arrows in Reaction 2.2 show the movement of pairs of electrons. These ar-
rows are used to depict reaction mechanisms and we will encounter many such dia-
grams throughout this book. One of the free pairs of electrons on the oxygen will con-
tribute to formation of a new O — H covalent bond between the oxygen atom of the
hydronium ion and a proton (H®) abstracted from a water molecule. An O — H cova-
lent bond is broken in this reaction and the electron pair from that bond remains asso-
ciated with the oxygen atom of the hydroxide ion.

Note that the atoms in the hydronium ion contain eleven positively charged pro-
tons (eight in the oxygen atom and three hydrogen protons) and ten negatively charged
electrons (a pair of electrons in the inner orbital of the oxygen atom, one free electron
pair associated with the oxygen atom, and three pairs in the covalent bonds). This results
in a net positive charge which is why we refer to it as an ion (cation). The positive charge
is usually depicted as if it were associated with the oxygen atom but, in fact, it is distrib-
uted partially over the hydrogen atoms as well. Similarly, the hydroxide ion (anion)
bears a net negative charge because it contains ten electrons whereas the nuclei of the
oxygen and hydrogen atoms have a total of only nine positively charged protons.

CHAPTER 2 Water

The density of water varies with the
temperature (Box 2.2) and so does the
ion product. The differences aren’t sig-
nificant in the temperature ranges that
we normally encounter in living cells,
so we assume that the value 10" 14
applies at all temperatures. (See
Problem 17 at the end of this chapter.)

The ionization reaction is a typical reversible reaction. The protonation and depro-
tonation reactions take place very quickly. Hydroxide ions have a short lifetime in water
and so do hydronium ions. Even water molecules themselves have only a transient exis-
tence. The average water molecule is thought to exist for about one millisecond (10 _3 s)
before losing a proton to become a hydroxide ion or gaining a proton to become a hy-
dronium ion. Note that the lifetime of a water molecule is still eight orders of magni-
tude (10 8 ) greater than the lifetime of a hydrogen bond.

Hydronium ( H 3 0©) ions are capable of donating a proton to another ion. Such
proton donors are referred to as acids according to the Bronsted-Lowry concept of
acids and bases. In order to simplify chemical equations we often represent the hydro-
nium ion as simply H© (free proton or hydrogen ion) to reflect the fact that it is a major
source of protons in biochemical reactions. The ionization of water can then be de-
picted as a simple dissociation of a proton from a single water molecule.

H 2 0 H© + OH© (2.3)

Reaction 2.3 is a convenient way to show the ionization of water but it does not re-
flect the true structure of the proton donor which is actually the hydronium ion. Reac-
tion 2.3 also obscures the fact that the ionization of water is actually a bimolecular reac-
tion involving two separate water molecules as shown in Reaction 2.2. Fortunately, the
dissociation of water is a reasonable approximation that does not affect our calculations
or our understanding of the properties of water. We will make use of this assumption in
the rest of the book.

Hydroxide ions can accept a proton and be converted back into water molecules.
Proton acceptors are called bases. Water can function as either an acid or a base as Reac-
tion 2.2 demonstrates.

The ionization of water can be analyzed quantitatively. Recall that the concentra-
tions of reactants and products in a reaction will eventually reach an equilibrium where
there is no net change in concentration. The ratio of these equilibrium concentrations
defines the equilibrium constant (K eq ). In the case of ionization of water,

Keq = [H « K eq [H 2 0] = [H@][OH©] (2.4)

The equilibrium constant for the ionization of water has been determined under stan-
dard conditions of pressure (1 atm) and temperature (25°C). Its value is 1.8 X 1(T 16 M. We
are interested in knowing the concentrations of protons and hydroxide ions in a solu-
tion of pure water since these ions participate in many biochemical reactions. These
values can be calculated from Equation 2.4 if we know the concentration of water
( [H 2 0]) at equilibrium. Pure water at 25°C has a concentration of approximately 55.5 M
(see Box 2.2). A very small percentage of water molecules will dissociate to form H©
and OH© when the ionization reaction reaches equilibrium. This will have a very small
effect on the final concentration of water molecules at equilibrium. We can simplify our
calculations by assuming that the concentration of water in Equation 2.4 is 55.5 M.
Substituting this value, and that of the equilibrium constant, gives

(1.8 X 10“ 16 M)(55.5 M) = 1.0 X 10“ 14 M 2 = [H©][OH e ] (2.5)

The product obtained by multiplying the proton and hydroxide ion concentrations
([H©] [OH©]) is called the ion product for water. This is a constant designated K w (the
ion product constant for water). At 25°C the value of K w is

K w = [H©][OH©] = 1.0 X 10“ 14 M 2 (2.6)

It is a fortunate coincidence that this is a nice round number rather than some awkward
fraction because it makes calculations of ion concentrations much easier. Pure water is

2.8 The pH Scale 43

electrically neutral, so its ionization produces an equal number of protons and hydroxide
ions [H©] = [OH] . In the case of pure water, Equation 2.6 can therefore be rewritten as

K w = [H©] 2 = 1.0 X 1(T 14 M 2 (2.7)

Taking the square root of the terms in Equation 2.7 gives

[H©] = 1.0X1 O -7 M (2.8)

Since [H©] = [OH©], the ionization of pure water produces 1CT 7 M H© and 1(T 7 M
OH©. Pure water and aqueous solutions that contain equal concentrations of H© and
OH© are said to be neutral. Of course, not all aqueous solutions have equal concentra-
tions of H© and OH©. When an acid is dissolved in water [H©] increases and the solu-
tion is described as acidic. Note that when an acid is dissolved in water the concentra-
tion of protons increases while the concentration of hydroxide ions decreases. This is
because the ion product constant for water (K w ) is unchanged (i.e., constant) and the
product of the concentrations of H© and OH© must always be 1.0 X 10 -14 M 2 under
standard conditions (Equation 2.5). Dissolving a base in water decreases [H©] and in-
creases [OH©] above 1.0 X 10 7 M producing a basic, or alkaline, solution.

2.8 The pH Scale

Many biochemical processes — including the transport of oxygen in the blood, the catal-
ysis of reactions by enzymes, and the generation of metabolic energy during respiration
or photosynthesis — are strongly affected by the concentration of protons. Although the
concentration of H® (or H 3 0©) in cells is small relative to the concentration of water,
the range of [H©] in aqueous solutions is enormous so it is convenient to use a loga-
rithmic quantity called pH as a measure of the concentration of H©. pH is defined as the
negative logarithm of the concentration of H©.

pH = -log[H@] = log^T_ (2.9)

In pure water [H©] = [OH©] = 1.0 X 10 -7 M (Equations 2.7 and 2.8). As men-
tioned earlier, pure water is said to be “neutral” with respect to total ionic charge since
the concentrations of the positively charged hydrogen ions and the negatively charged
hydroxide ions are equal. Neutral solutions have a pH value of 7.0 (the negative value of
log 10 -7 is 7.0). Acidic solutions have an excess of H© due to the presence of dissolved
solute that supplies H© ions. In a solution of 0.01 M HC1, for example, the concentra-
tion of H© is 0.01 M (10 -2 M) because HC1 dissociates completely to H© and Cl©. The
pH of such a solution is -log 10 -2 = 2.0. Thus, the higher the concentration of H©, the
lower the pH of the solution. The pH scale is logarithmic, so a change in pH of one unit
corresponds to a 10-fold change in the concentration of H©.

Aqueous solutions can also contain fewer H© ions than pure water resulting in a
pH above 7. In a solution of 0.01 M NaOH, for example, the concentration of OH© is
0.01 M (10 -2 M) because NaOH, like HC1, is 100% dissociated in water. The H© ions
derived from the ionization of water will combine with the hydroxide ions from NaOH
to re-form water molecules. This affects the equilibrium for the ionization of water
(Reaction 2.3). The resulting solution is very basic because of the low concentration of
protons. The actual pH can be determined from the ion product of water, K w (Equa-
tion 2.6), by substituting the concentration of hydroxide ions. Since the product of the
OH© and H© concentrations is 10 -14 M it follows that the H© concentration in a solution
of 1(T 2 M OH© is 10 -12 M. The pH of the solution is 12. Table 2.3 shows this relationship
between pH and the concentrations of H© and OH©.

Basic solutions have pH values greater than 7.0 and acidic solutions have lower
pH values. Figure 2.16 illustrates the pH values of various common solutions.

Figure 2.16 ►

pH values for various fluids at 25°C. Lower values correspond to acidic fluids; higher values corre-
spond to basic fluids.

Table 2.3 Relation of [H©] and [0H @ ] to pH

[H®]

(M)

[OH©]

(M)

IO -14

10 _1

IO" 13

10“ 2

io- 12

1(T 3

IO -11

10 4

IQ- 10

10 5

10 9

10“ 6

10“ 8

io- 7

10“ 8

10“ 6

IO" 9

10 5

10~ 10

IO 4

10 H1

IO 3

10- 12

10“ 2

io- 13

10 _1

IO -14

Sodium

hydroxide (1 M)

Ammonia (1 M)

Milk of Magnesia

-M

l/l

Human

pancreatic

juice

Human

blood

plasma

Cow's milk

Coffee (black)

Tomato

juice

Wine

Lemon juice
Human
stomach
secretions

Hydrochloric
acid (1 M)

44 CHAPTER 2 Water

tOO Slr ips

pH indicator attipa ntm-btoilinB

pH pH o - 14

▲ pH strips. The approximate pH of solutions
can be determined in the lab by placing a
drop on a pH strip. Various indicators are
bound to a matrix that is affixed to a plastic
strip. The indicators change color at different
concentrations of H®, and the combination of
various colors gives a more or less accurate
reading of the pH. The strips shown here cover
all pH readings from 0 to 14 but other pH
strips can be used to cover narrower ranges.

KEY CONCEPT

pH is the negative logarithm of the proton
(H©) concentration.

BOX 2.4 THE LITTLE “p” IN pH

The term pH was first used in 1909 by S 0 ren
Peter Lauritz Sorensen, director of the Carls-
berg Laboratories in Denmark. Sorensen never
mentioned what the little “p” stood for (the ££ H”
is obviously hydrogen). Many years later, some
of the scientists who write chemistry textbooks
began to associate the little “p” with the words
power or potential. This association, as it turns
out, is based on a rather tenuous connection in
some of Sorensens early papers. A recent inves-
tigation of the historical records by Jens G.
Noby suggests that the little “p” was an arbitrary
choice based on Sorensen’s use of p and q to
stand for unknown variables in much the same
way that we might use x and y today.

No matter what the historical origin, it’s
important to remember that the symbol pH
now stands for the negative logarithm of the
hydrogen ion concentration.

▲ Spren Peter Lauritz Sprensen
( 1868 - 1939 )

Accurate measurements of pH are routinely made using a pH meter, an instrument
that incorporates a selectively permeable glass electrode that is sensitive to [H©].
Measurement of pH sometimes facilitates the diagnosis of disease. The normal pH of
human blood is 7.4 — frequently referred to as physiological pH. The blood of pa-
tients suffering from certain diseases, such as diabetes, can have a lower pH, a condi-
tion called acidosis. The condition in which the pH of the blood is higher than 7.4,
called alkalosis, can result from persistent, prolonged vomiting (loss of hydrochloric
acid from the stomach) or from hyperventilation (excessive loss of carbonic acid as
carbon dioxide).

KEY CONCEPT

Weak acids and weak bases are
compounds that only partially dissociate
in water.

2.9 Acid Dissociation Constants of Weak Acids

Acids and bases that dissociate completely in water, such as hydrochloric acid and
sodium hydroxide, are called strong acids and strong bases. Many other acids and bases,
such as the amino acids from which proteins are made and the purines and pyrimidines
from DNA and RNA, do not dissociate completely in water. These substances are
known as weak acids and weak bases.

In order to understand the relationship between acids and bases let us consider the
dissociation of HC1 in water. Recall from Section 2.7 that we define an acid as a mole-
cule that can donate a proton and a base as a proton acceptor. Acids and bases always
come in pairs since for every proton donor there must be a proton acceptor. Both sides
of the dissociation reaction will contain an acid and a base. Thus, the equilibrium reac-
tion for the complete dissociation of HC1 is

HCI + H 2 0 Cl 0 + H 3 0® (2.10)

acid base base acid

HCI is an acid because it can donate a proton. In this case, the proton acceptor is
water which is the base in this equilibrium reaction. On the other side of the equilib-
rium are Cl© and the hydronium ion, H 3 0©. The chloride ion is the base that corre-
sponds to HCI after it has given up its proton. Cl© is called the conjugate base of HCI
which indicates that it is a base (i.e., can accept a proton) and is part of an acid-base
pair (i.e., HC1/C1©). Similarly, H 3 0© is the acid on the right-hand side of the equi-
librium because it can donate a proton. H 3 0© is the conjugate acid of H 2 0. Every base

2.9 Acid Dissociation Constants of Weak Acids

has a corresponding conjugate acid and every acid has a corresponding conjugate
base. Thus, HC1 is the conjugate acid of Cl® and H 2 0 is the conjugate base of H 3 0®.
Note that H 2 0 is the conjugate acid of OH® if we are referring to the H 2 0/0H®
acid-base pair.

In most cases throughout this book we will simplify reactions by ignoring the con-
tribution of water and representing the hydronium ion as a simple proton.

HCI H© + Cl© ( 2 . 11 )

This is a standard convention in biochemistry but, on the surface, it seems to violate the
rule that both sides of the equilibrium reaction should contain a proton donor and a
proton acceptor. Students should keep in mind that in such reactions the contributions
of water molecules as proton acceptors and hydronium ions as the true proton donors
are implied. In almost all cases we can safely ignore the contribution of water. This is the
same principle that we applied to the reaction for the dissociation of water (Section 2.7)
which we simplified by ignoring the contribution of one of the water molecules.

The reason why HC1 is such a strong acid is because the equilibrium shown in Re-
action 2.11 is shifted so far to the right that HC1 is completely dissociated in water. In
other words, HC1 has a strong tendency to donate a proton when dissolved in water.
This also means that the conjugate base, Cl®, is a very weak base because it will rarely
accept a proton.

Acetic acid is the weak acid present in vinegar. The equilibrium reaction for the
ionization of acetic acid is

KEY CONCEPT

The contribution of water is implied in
most acid/base dissociation reactions.

CH 3 COOH H© + CH 3 COO© ( 2 . 12 )

Acetic acid Acetate anion

(weak acid) (conjugate base)

We have left out the contribution of water molecules in order to simplify the reaction.
We see that the acetate ion is the conjugate base of acetic acid. (We can also refer to acetic
acid as the conjugate acid of the acetate ion.)

The equilibrium constant for the dissociation of a proton from an acid in water is
called the acid dissociation constant, K a . When the reaction reaches equilibrium, which
happens very rapidly, the acid dissociation constant is equal to the concentration of the
products divided by the concentration of the reactants. For Reaction 2.12 the acid dis-
sociation constant is

[H©][CH 3 COO©]
a [CH 3 COOH]

The K a value for acetic acid at 25°C is 1.76 x 10 -5 M. Because K a values are numeri-
cally small and inconvenient in calculations it is useful to place them on a logarithmic
scale. The parameter p K a is defined by analogy with pH.

P K a = -log K a = log - 7 - (2.14)

K a

A pH value is a measure of the acidity of a solution and a p K a value is a measure of
the acid strength of a particular compound. The p K a of acetic acid is 4.8.

When dealing with bases we need to consider their protonated forms in order to
use Equation 2.13. These conjugate acids are very weak acids. In order to simplify calcu-
lations and make easy comparisons we measure the equilibrium constant (K a ) for the
dissociation of a proton from the conjugate acid of a weak base. For example, the am-
monium ion (NH 4 ®) can dissociate to form the base ammonia (NH 3 ) and H®.

NH 4 ® NH 3 + H®

(2.15)

The acid dissociation constant (K a ) for this equilibrium is a measure of the strength of
the base (ammonia, NH 3 ) in aqueous solution. The K a values for several common sub-
stances are listed in Table 2.4.

CHAPTER 2 Water

Table 2.4 Dissociation constants and pK a values of weak acids in aqueous
solutions at 25°C

Acid

K a(M)

pK a

HCOOH (Formic acid)

1.77 X 10 “ 4

3.8

CH 3 COOH (Acetic acid)

1.76 X 1(T 5

4.8

CH 3 CHOHCOOH (Lactic acid)

1.37 X 10 “ 4

3.9

H 3 PO 4 (Phosphoric acid)

7.52 X 10 “ 3

2.2

H 2 P0 4 ® (Dihydrogen phosphate ion)

HPO 4 (Monohydrogen phosphate ion)

6.23 X 1(T 8

7.2

2.20 X 10 “ 13

12.7

H 2 C0 3 (Carbonic acid)

4.30 X 10 “ 7

6.4

HCO 3 0 (Bicarbonate ion)

5.61 X 10~ n

10.2

NH 4 © (Ammonium ion)

5.62 X 10 “ 10

9.2

CH 3 NH 3 © (Methylammonium ion)

2.70 X 10 -11

10.7

From Equation 2.13 we see that the FC a for acetic acid is related to the concentra-
tion of H® and to the ratio of the concentrations of the acetate ion and undissociated
acetic acid. If we represent the conjugate acid as HA and the conjugate base as A® then
taking the logarithm of such equations gives the general equation for any acid-base
pair.

HA H© + A© log K a = log

[H©][A 0 ]

[HA]

(2.16)

Since log(xy) = log x + logy, Equation 2.16 can be rewritten as

[A©]

log K a = log[H©] + log

[HA]

(2.17)

Rearranging Equation 2.17 gives

-log[H©] = -log K a + log

[A Q ]

[HA]

(2.18)

KEY CONCEPT

The pH of a solution of a weak acid or
base at equilibrium can be calculated by
combining the p K a of the ionization
reaction and the final concentrations of
the proton acceptor and proton donor
species.

The negative logarithms in Equation 2.18 have already been defined as pH and p K a
(Equations 2.9 and 2.14, respectively). Thus,

, [A e ]

pH - pK ‘ + Io 3 IhaI

(2.19)

PH = p K a

+ log

[Proton acceptor]
[Proton donor]

( 2 . 20 )

Equation 2.20 is one version of the Henderson-Hasselbalch equation. It defines the
pH of a solution in terms of the pFC a of the weak acid form of the acid-base pair and
the logarithm of the ratio of concentrations of the dissociated species (conjugate base)
to the protonated species (weak acid). Note that the greater the concentration of the
proton acceptor (conjugate base) relative to that of the proton donor (weak acid),
the lower the concentration of H® and the higher the pH. (Remember that pH is the
negative log of H® concentration. A high concentration of H® means low pH.) This

2.9 Acid Dissociation Constants of Weak Acids 47

makes intuitive sense since the concentration of A© is identical to the concentration of
H© in simple dissociation reactions. If more HA dissociates the concentration of A©
will be higher and so will the concentration of H©. When the concentrations of a weak
acid and its conjugate base are exactly the same the pH of the solution is equal to the
p K a of the acid (since the ratio of concentrations equals 1.0, and the logarithm of 1.0
equals zero).

The Henderson-Hasselbalch equation is used to determine the final pH of a weak
acid solution once the dissociation reaction reaches equilibrium as illustrated in Sample
Calculation 2.1 for acetic acid. These calculations are more complicated than those in-
volving strong acids such as HC1. As noted in Section 2.8, the pH of an HC1 solution is
easily determined from the amount of HC1 that is present since the final concentration
of H© is equal to the initial concentration of HC1 when the solution is made up. In con-
trast, weak acids are only partially dissociated in water so it makes sense that the pH de-
pends on the acid dissociation constant. The pH decreases (more H©) as more weak
acid is added to water but the increase in H© is not linear with initial HA concentra-
tion. This is because the numerator in Equation 2.16 is the product of the H© and A©
concentrations.

The Henderson-Hasselbalch equation applies to other acid-base combinations as
well and not just to those involving weak acids. When dealing with a weak base, for ex-
ample, the numerator and denominator of Equation 2.20 become [weak base] and
[conjugate acid], respectively. The important point to remember is that the equation
refers to the concentration of the proton acceptor divided by the concentration of the
proton donor.

The pK a values of weak acids are determined by titration. Figure 2.17 shows the
titration curve for acetic acid. In this example, a solution of acetic acid is titrated by
adding small aliquots of a strong base of known concentration. The pH of the solution
is measured and plotted versus the number of molar equivalents of strong base added
during the titration. Note that since acetic acid has only one ionizable group (its car-
boxyl group) only one equivalent of a strong base is needed to completely titrate acetic
acid to its conjugate base, the acetate anion. When the acid has been titrated with one-
half an equivalent of base the concentration of undissociated acetic acid exactly equals
the concentration of the acetate anion. The resulting pH, 4.8, is thus the experimentally
determined p for acetic acid.

Constructing an ideal titration curve is a useful exercise for reinforcing the rela-
tionship between pH and the ionization state of a weak acid. You can use the Hender-
son-Hasselbalch equation to calculate the pH that results from adding increasing amounts
of a strong base such as NaOH to a weak acid such as the imidazolium ion p K a = 7.0.
Adding base converts the imidazolium ion to its conjugate base, imidazole (Figure 2.18).
The shape of the titration curve is easy to visualize if you calculate the pH when the
ratio of conjugate base to acid is 0.01, 0.1, 1, 10, and 100. Calculate pH values at other
ratios until you are satisfied that the curve is relatively flat near the midpoint and
steeper at the ends.

Similarly shaped titration curves can be obtained for each of the five monoprotic
acids (acids having only one ionizable group) listed in Table 2.4. All would exhibit the
same general shape as Figure 2.17 but the inflection point representing the midpoint of
titration (one-half an equivalent titrated) would fall lower on the pH scale for a
stronger acid (such as formic acid or lactic acid) and higher for a weaker acid (such as
ammonium ion or methylammonium ion).

Titration curves of weak acids illustrate a second important use of the Henderson-
Hasselbalch equation. In this case, the final pH is the result of mixing the weak acid
(HA) and a strong base (OH©). The base combines with H© ions to form water mole-
cules, H 2 0. This reduces the concentration of H© and raises the pH. As the titration of
the weak acid proceeds it dissociates in order to restore its equilibrium with OH© and
H 2 0. The net result is that the final concentration of A© is much higher, and the con-
centration of HA is much lower, than when we are dealing with the simple case where
the pH is determined only by the dissociation of the weak acid in water (i.e., a solution
of HA in H 2 0).

▲ Figure 2.17

Titration of acetic acid (CH 3 C00H) with aque-
ous base (OH®). There is an inflection point
(a point of minimum slope) at the midpoint
of the titration, when 0.5 equivalent of base
has been added to the solution of acetic
acid. This is the point at which
[CH 3 COOH] = [CH 3 C00 e ] and pH = pK a .
The p K a of acetic acid is thus 4.8. At the
endpoint, all the molecules of acetic acid
have been titrated to the conjugate base,
acetate.

— H

Imidazolium ion

H ©

P K a = 7.0

Imidazole

▲ Figure 2.18

Titration of the imidazolium ion.

48 CHAPTER 2 Water

Figure 2.19 ►

Titration curve for H 3 P0 4 . Three inflection
points (at 0.5, 1.5, and 2.5 equivalents of
strong base added) correspond to the three
p K a values for phosphoric acid (2.2, 7.2,
and 12.7).

▲ Cola beverages contain phosphoric acid
in order to make the drink more acidic. The
concentration of phosphoric acid is about
1 mM. This concentration should make the
pH about 3 in the absence of any other
ingredients that may contribute to acidity.

Third midpoint

[hpo 4 ®] = [po 4 ®]

Phosphoric acid (H 3 PO 4 ) is a polyprotic acid. It contains three different hydrogen
atoms that can dissociate to form H© ions and corresponding conjugate bases with one,
two, or three negative charges. The dissociation of the first proton occurs readily and is
associated with a large acid dissociation constant of 7.53 x 10 -3 M and a pl<f a of 2.2 in
aqueous solution. The dissociations of the second and third protons occur progressively
less readily because they have to dissociate from a molecule that is already negatively
charged.

Phosphoric acid requires three equivalents of strong base for complete titration
and three p FC a values are evident from its titration curve (Figure 2.19). The three pFC a
values reflect the three equilibrium constants and thus the existence of four possible
ionic species (conjugate acids and bases) of inorganic phosphate. At physiological pH
(7.4) the predominant species of inorganic phosphate are H 2 P0 4 © and HP0 4 ©. At
pH 7.2 these two species exist in equal concentrations. The concentrations of H 3 P0 4
and P0 4 © are so low at pH 7.4 that they can be ignored. This is generally the case for a
minor species when the pH is more than two units away from its p K a .

HO— P— OH

PKi

2.2

II 0

HO— P—O

p k 2

7.2

o— p— o

H©

H ©

p/C 3

12.7

( 2 . 21 )

O— P— o

I©

H ©

Many biologically important acids and bases, including the amino acids described
in Chapter 3, have two or more ionizable groups. The number of p K a values for such
substances is equal to the number of ionizable groups. The p K a values can be experi-
mentally determined by titration.

2.9 Acid Dissociation Constants of Weak Acids

Sample Calculation 2.1 CALCULATING THE pH OF WEAK ACID

SOLUTIONS

Q: What is the pH of a solution of 0.1 M acetic acid?

A: The acid dissociation constant of acetic acid is 1.76 X 10 -5 M. Acetic acid disso-
ciates in water to form acetate and H® . We need to determine [H® ] when the reaction
reaches equilibrium.

Let the final H® concentration be represented by the unknown quantity x. At equi-
librium the concentration of acetate ion will also be x and the final concentration of
acetic acid will be [0.1 M — x]. Thus,

, 7 , x 1(r s . [H e ][CH 3 C00 9 ] _

[CHjCOOH] (0.1 - x)

rearranging gives

1 .76 X 1 0“ 6 - 1 .76 X 1 0“ 5 x = x 2
x 2 + 1 .76 X 1 0“ 5 x - 1 .76 X 1 0“ 6 = 0

This equation is a typical quadratic equation of the form ax 2 + bx + c = 0, where
a = 1, b = 1.76 X 10 -5 , and c = —1.76 X 10 -6 . Solve for x using the standard
formula

-b ± V(b 2 - 4oc)

-1.76 X 10“ 5 ± V((1 .76 X 10“ 5 ) 2 - 4(1.76 X 10“ 6 ))

- 2
x = 0.001 32 or -0.001 35 (reject the negative answer)

The hydrogen ion concentration is 0.00132 M and the pH is

pH = -log[H®] = -log(0.001 32) = -(-2.88) = 2.9

Note that the contribution of hydrogen ions from the dissociation of water 110 7 2 is
several orders of magnitude lower than the concentration of hydrogen ions from
acetic acid. It is standard practice to ignore the ionization of water in most calcula-
tions as long as the initial concentration of weak acid is greater than 0.001 M.

The amount of acetic acid that dissociates to form H® and CH 3 COO® is 0.0013 M
when the initial concentration is 0.1 M. This means that only 1.3% of the acetic acid
molecules dissociate and the final concentration of acetic acid l[CH 3 COOH]2 is
98.7% of the initial concentration. In general, the percent dissociation of dilute
solutions of weak acids is less than 10% and it is a reasonable approximation to
assume that the final concentration of the acid form is the same as its initial concen-
tration. This approximation has very little effect on the calculated pH and it has the
advantage of avoiding quadratic equations.

Assuming that the concentration of CH3COOH at equilibrium is 0.1 M and the con-
centration of H® is x,

x 2

K a = 1.76 X 1CT 5 = — x = 1 .33 X 1 0 -3
pH = — log( 1 .33 X 1CT 3 ) = 2.88 = 2.9

CH 2 OH

hoh 2 c — c — NH,

ch 2 oh

▲ Tris buffers. Tris, or tris (hydroxymethyl)
aminomethane, is a common buffer in
biochemistry labs. Its p K a of 8.06 makes
it ideal for preparation of buffers in the
physiological range.

50 CHAPTER 2 Water

2.10 Buffered Solutions Resist Changes in pH

▲ Figure 2.20

Buffer range of acetic acid. For CH 3 COOH +
CH 3 COO 0 the p K a is 4.8 and the most ef-
fective buffer range is from pH 3.8 to pH
5.8.

If the pH of a solution remains nearly constant when small amounts of strong acid or
strong base are added the solution is said to be buffered. The ability of a solution to resist
changes in pH is known as its buffer capacity. Inspection of the titration curves of acetic
acid (Figure 2.17) and phosphoric acid (Figure 2.19) reveals that the most effective
buffering, indicated by the region of minimum slope on the curve, occurs when the
concentrations of a weak acid and its conjugate base are equal — in other words, when
the pH equals the p K a . The effective range of buffering by a mixture of a weak acid and
its conjugate base is usually considered to be from one pH unit below to one pH unit
above the p K a .

Most in vitro biochemical experiments involving purified molecules, cell extracts,
or intact cells are performed in the presence of a suitable buffer to ensure a stable pH. A
number of synthetic compounds with a variety of p K a values are often used to prepare
buffered solutions but naturally occurring compounds can also be used as buffers. For
example, mixtures of acetic acid and sodium acetate (p K a = 4.8) can be used for the pH
range from 4 to 6 (Figure 2.20) and mixtures of KH 2 P0 4 and K 2 HP0 4 (pFC a = 7.2) can be
used in the range from 6 to 8. The amino acid glycine (p K a = 9.8) is often used in the
range from 9 to 11.

When preparing buffers the acid solution (e.g., acetic acid) supplies the protons
and some of the protons are taken up by combining with the conjugate base (e.g., ac-
etate). The conjugate base is added as a solution of a salt (e.g., sodium acetate). The salt
dissociates completely in solution providing free conjugate base and no protons.
Sample Calculation 2.2 illustrates one way to prepare a buffer solution.

Sample Calculation 2.2 BUFFER PREPARATION

Q: Acetic acid has a p K a of 4.8. How many milliliters of 0.1 M acetic acid and 0.1 M
sodium acetate are required to prepare 1 liter of 0.1 M buffer solution having a pH
of 5.8?

A: Substitute the values for the p K a and the desired pH into the Henderson-Hassel-
balch equation (Equation 2.20).

5.8

4.8 +

log

[Acetate]
[Acetic acid]

Solve for the ratio of acetate to acetic acid.

log

[Acetate]
[Acetic acid]

= 5.8 - 4.8 = 1.0

[Acetate] = 1 0 [Acetic acid]

For each volume of acetic acid, 10 volumes of acetate must be added (making a total
of 1 1 volumes of the two ionic species). Multiply the proportion of each component
by the desired volume.

Acetic acid needed: A x 1000 ml = 91 ml

Acetate needed: — x 1000 ml = 909 ml

Note that when the ratio of [conjugate base] to [conjugate acid] is 10:1, the pH is ex-
actly one unit above the p/v a . If the ratio were. 1:10, the pH would be one unit below
the p K a .

2.10 Buffered Solutions Resist Changes in pH 51

► Figure 2.21

Percentages of carbonic acid and its conjugate
base as a function of pH. In an aqueous
solution at pH 7.4 (the pH of blood) the
concentrations of carbonic acid (H 2 C0 3 ) and
bicarbonate (HCO 3 0 ) are substantial, but
the concentration of carbonate (C0 3 ©) is
negligible.

An excellent example of buffer capacity is found in the blood plasma of mammals,
which has a remarkably constant pH. Consider the results of an experiment that compares
the addition of an aliquot of strong acid to a volume of blood plasma with a similar addi-
tion of strong acid to either physiological saline (0.15 M NaCl) or water. When 1 milliliter
of 10 M HC1 (hydrochloric acid) is added to 1 liter of physiological saline or water that
is initially at pH 7.0 the pH is lowered to 2.0 (in other words, [H©] from HC1 is diluted
to 10 -2 M). However, when 1 milliliter of 10 M HC1 is added to 1 liter of human blood
plasma at pH 7.4 the pH is lowered to only 7.2 — impressive evidence for the effective-
ness of physiological buffering.

The pH of blood is primarily regulated by the carbon dioxide-carbonic acid-bicar-
bonate buffer system. A plot of the percentages of carbonic acid (H 2 C0 3 ) and its conju-
gate base as a function of pH is shown in Figure 2.21. Note that the major components
at pH 7.4 are carbonic acid and the bicarbonate anion (HC0 3 ©).

The buffer capacity of blood depends on equilibria between gaseous carbon diox-
ide (which is present in the air spaces of the lungs), aqueous carbon dioxide (which is
produced by respiring tissues and dissolved in blood), carbonic acid, and bicarbonate.
As shown in Figure 2.21, the equilibrium between bicarbonate and its conjugate base,
carbonate (C0 3 ©), does not contribute significantly to the buffer capacity of blood be-
cause the p K a of bicarbonate is 10.2 — too far from physiological pH to have an effect on
the buffering of blood.

The first of the three relevant equilibria of the carbon dioxide-carbonic acid-bicar-
bonate buffer system is the dissociation of carbonic acid to bicarbonate.

H 2 C0 3 H© + HCO 3 0 ( 2 . 22 )

This equilibrium is affected by a second equilibrium in which dissolved carbon dioxide
is in equilibrium with its hydrated form, carbonic acid.

C0 2 (aqueous) + H 2 0 H 2 C0 3 (2.23)

These two reactions can be combined into a single equilibrium reaction where the acid
is represented as C0 2 dissolved in water:

C0 2 (aqueous) + H 2 Q H© + HC0 3 © (2.24)

Aqueous phase
of blood cells
passing through
capillaries
in lung

hco 3 °

>H©

h 2 c

:o 3

h 2 o^

^-HzO

C0 2

(aqueous)

C0 2

(gaseous)

Air space
in lung

The p K a of the acid is 6.4.

Finally, C0 2 (gaseous) is in equilibrium with C0 2 (aqueous).

C0 2 (gaseous) C0 2 (aqueous) (2.25)

The regulation of the pH of blood afforded by these three equilibria is shown
schematically in Figure 2.22. When the pH of blood falls due to a metabolic process that
produces excess H© the concentration of H 2 C0 3 increases momentarily but H 2 C0 3

▲ Figure 2.22

Regulation of the pH of blood in mammals. The

pH of blood is controlled by the ratio of
[HC0 3 ®] topC0 2 in the air spaces of the
lungs. When the pH of blood decreases due
to excess H®, pC0 2 increases in the lungs,
restoring the equilibrium. When the concen-
tration of HCO 3 0 rises because the pH of
blood increases, C0 2 (gaseous) dissolves in
the blood, again restoring the equilibrium.

52 CHAPTER 2 Water

rapidly loses water to form dissolved C0 2 (aqueous) which enters the gaseous phase in
the lungs and is expired as C0 2 (gaseous). An increase in the partial pressure of C0 2
( pC0 2 ) in the air expired from the lungs thus compensates for the increased hydrogen
ions. Conversely, when the pH of the blood rises the concentration of HC0 3 ® increases
transiently but the pH is rapidly restored as the breathing rate changes and the C0 2
(gaseous) in the lungs is converted to C0 2 (aqueous) and then to H 2 C0 3 in the capillar-
ies of the lungs. Again, the equilibrium of the blood buffer system is rapidly restored by
changing the partial pressure of C0 2 in the lungs.

Within cells, both proteins and inorganic phosphate contribute to intracellular
buffering. Hemoglobin is the strongest buffer in blood cells other than the carbon diox-
ide-carbonic acid-bicarbonate buffer. As mentioned earlier, the major species of inor-
ganic phosphate present at physiological pH are H 2 P0 4 ® and HP0 4 © reflecting the
second p K a (p K 2 ) value for phosphoric acid, 7.2.

Summary

1. The water molecule has a permanent dipole because of the un-
even distribution of charge in O — H bonds and their angled
arrangement.

2. Water molecules can form hydrogen bonds with each other. Hy-
drogen bonding contributes to the high specific heat and heat of
vaporization of water.

3. Because it is polar, water can dissolve ions. Water molecules form
a solvation sphere around each dissolved ion. Organic molecules
may be soluble in water if they contain ionic or polar functional
groups that can form hydrogen bonds with water molecules.

4. The hydrophobic effect is the exclusion of nonpolar substances by
water molecules. Detergents, which contain both hydrophobic and
hydrophilic portions, form micelles when suspended in water; these
micelles can trap insoluble substances in a hydrophobic interior.
Chaotropes enhance the solubility of nonpolar compounds in water.

5. The major noncovalent interactions that determine the structure and
function of biomolecules are electrostatic interactions and hydropho-
bic interactions. Electrostatic interactions include charge-charge
interactions, hydrogen bonds, and van der Waals forces.

6. Under cellular conditions, macromolecules do not spontaneously
hydrolyze, despite the presence of high concentrations of water.
Specific enzymes catalyze their hydrolysis, and other enzymes cat-
alyze their energy- requiring biosynthesis.

7. At 25°C, the product of the proton concentration ( [H®] ) and the
hydroxide concentration ([OH®]) is 1.0 x 1CT 14 M 2 , a constant
designated K w (the ion-product constant for water). Pure water
ionizes to produce 1(T 7 M H® and 1(T 7 M OH®.

8. The acidity or basicity of an aqueous solution depends on the
concentration of H® and is described by a pH value, where pH is
the negative logarithm of the hydrogen ion concentration.

9. The strength of a weak acid is indicated by its pK a value. The
Henderson-Hasselbalch equation defines the pH of a solution of
weak acid in terms of the p K a and the concentrations of the weak
acid and its conjugate base.

10. Buffered solutions resist changes in pH. In human blood, a con-
stant pH of 7.4 is maintained by the carbon dioxide-carbonic
acid-bicarbonate buffer system.

Problems

1. The side chains of some amino acids possess functional groups
that readily form hydrogen bonds in aqueous solution. Draw the
hydrogen bonds likely to form between water and the following
amino acid side chains:

(a)

(b)

(c)

ch 2 oh

CH 2 C(0)NH 2

— CH-

N =

V-N — H

2. State whether each of the following compounds is polar, whether
it is amphipathic, and whether it readily dissolves in water.

(a) HO — CH 2 — CH — CH 2 — OH

Glycerol

(b) ch 3 ich 2 2 14 — ch 2 — opo 3

Hexadecanyl phosphate

Laurate

(d) h 3 n — ch 2 — coo g

Glycine

3. Osmotic lysis is a gentle method of breaking open animal cells to
free intracellular proteins. In this technique, cells are suspended
in a solution that has a total molar concentration of solutes much
less than that found naturally inside cells. Explain why this tech-
nique might cause cells to burst.

4. Each of the following molecules is dissolved in buffered solutions
of: (a) pH = 2 and (b) pH = 11. For each molecule, indicate the
solution in which the charged species will predominate. (Assume
that the added molecules do not appreciably change the pH of the
solution.)

(a) Phenyl lactic acid pK a = 4

CH 2 CH(OH)COOH

Problems 53

(b) Imidazole pK a = 1

ii ©

ch 3 occh 2 ch 2 ch 2 — nh 3

(d) Phenyl salicylate pK a = 9.6

5. Use Figure 2. 16 to determine the concentration of H® and OH® in:

(a) tomato juice

(b) human blood plasma

6. The interaction between two (or more) molecules in solution can
be mediated by specific hydrogen bond interactions. Phorbol es-
ters can act as a tumor promoter by binding to certain amino
acids that are part of the enzyme protein kinase C (PKC). Draw
the hydrogen bonds expected in the complex formed between the
tumor promoter phorbol and the glycine portion of PKC:
— NHCH 2 C(0)—

The nitrogen atom of MOPS can be protonated (pK a = 7.2). The
carboxyl group of SHS can be ionized (pK a = 5.5). Calculate the
ratio of basic to acidic species for each buffer at pH 6.5.

10 . Many phosphorylated sugars (phosphate esters of sugars) are
metabolic intermediates. The two ionizable — OH groups of the
phosphate group of the monophosphate ester of ribose (ribose 5-
phosphate) have pK a values of 1.2 and 6.6. The fully protonated
form of a-D-ribose 5 -phosphate has the structure shown below.

(a) Draw, in order, the ionic species formed upon titration of
this phosphorylated sugar from pH 0.0 to pH 10.0.

(b) Sketch the titration curve for ribose 5-phosphate.

11. Normally, gaseous C0 2 is efficiently expired in the lungs. Under
certain conditions, such as obstructive lung disease or emphy-
sema, expiration is impaired. The resulting excess of C0 2 in the
body may lead to respiratory acidosis, a condition in which excess
acid accumulates in bodily fluids. How does excess C0 2 lead to
respiratory acidosis?

12. Organic compounds in the diets of animals are a source of basic
ions and may help combat nonrespiratory types of acidosis. Many
fruits and vegetables contain salts of organic acids that can be me-
tabolized, as shown below for sodium lactate. Explain how the
salts of dietary acids may help alleviate metabolic acidosis.

Phorbol

7 . What is the concentration of a lactic acid buffer (pK a = 3.9) that
contains 0.25 M CH 3 CH(OH)COOH and 0.15 M CH 3 CH(OH)
COO®? What is the pH of this buffer?

8. You are instructed to prepare 100 ml of a 0.02 M sodium phos-
phate buffer, pH 7.2, by mixing 50 ml of solution A (0.02M
Na 2 HP0 4 ) and 50 ml of solution B (0.02 M NaH 2 P0 4 ). Refer to
Table 2.4 to explain why this procedure provides an effective
buffer at the desired pH and concentration.

9. What are the effective buffering ranges of MOPS (3-(N-mor-
pholino)propanesulfonic acid) and SHS (sodium hydrogen succi-
nate)?

MOPS

HOOC — CH 2 — CH 2 — COO® Na®
SHS

CH 3 — CH — COO®Na® + 30 2 »

Na® + 2 C0 2 + HC0 3 ® + 2H 2 0

13. Absorption of food in the stomach and intestine depends on the
ability of molecules to penetrate the cell membranes and pass into
the bloodstream. Because hydrophobic molecules are more likely
to be absorbed than hydrophilic or charged molecules, the ab-
sorption of orally administered drugs may depend on their pK a
values and the pH in the digestive organs. Aspirin (acetylsalicylic
acid) has an ionizable carboxyl group (pK a = 3.5). Calculate the
percentage of the protonated form of aspirin available for absorp-
tion in the stomach (pH = 2.0) and in the intestine (pH = 5.0).

Aspirin

14 . What percent of glycinamide, ®H 3 NCH 2 CONH 2 (pK a = 8.20) is
unprotonated at (a) pH 7.5, (b) pH 8.2, and (c) pH 9.0?

54 CHAPTER 2 Water

15 . Refer to the following table and titration curve to determine which
compound from the table is illustrated by the titration curve.

Compound

pKl

P«2

pK 3

Phosphoric acid

2.15

7.20

12.15

Acetic acid

4.76

Succinic acid

4.21

5.64

Boric acid

9.24

12.74

Glycine

2.40

9.80

16 . Predict which of the following substances are soluble in water.

CH 2 OH

about 4.0 x 10 13 . What is the actual neutral pH for extremophiles
living at 0°C and 100°C?

18. What is the approximate pH of a solution of 6 M HC1? Why doesn’t
the scale in Figure 2.16 accommodate the pH of such a solution?

Selected Readings

Water

Chaplin, M. F. (2001). Water, its importance
to life. Biochem. and Mol. Biol. Education
29:54-59.

Dix, J. A. and Verkman, A. S. (2008). Crowding ef-
fects on diffusion in solutions and cells. Annu. Rev.
Biophys. 37:247-263.

Stillinger, F. H. (1980). Water revisited. Science
209:451-457.

Verkman, A. S. (2001). Solute and macromolecular
diffusion in cellular aqueous compartments.

Trends Biochem Sci. 27:27-33.

Noncovalent Interactions

Fersht, A. R. (1987). The hydrogen bond in molec-
ular recognition. Trends Biochem. Sci. 12:301-304.

Frieden, E. (1975). Non-covalent interactions.

J. Chem. Educ. 52:754-761.

Tanford, C. (1980). The Hydrophobic Effect:
Formation of Micelles and Biological Membranes,
2nd ed. (New York: John Wiley & Sons).

Biochemical Calculations

Montgomery, R., and Swenson, C. A. (1976).
Quantitative Problems in Biochemical Sciences, 2nd
ed. (San Francisco: W. H. Freeman).

Segel, I. H. (1976). Biochemical Calculations:
How to Solve Mathematical Problems in General
Biochemistry, 2nd ed. (New York: John Wiley
& Sons).

pH and Buffers

Stoll, V. S., and Blanchard, J. S. (1990). Buffers:
principles and practice. Methods Enzymol.
182:24-38.

Norby, J. G. (2000). The origin and
meaning of the little p in pH.

Trends Biochem. Sci. 25:36-3 7.

Amino Acids and the Primary
Structures of Proteins

T he relationship between structure and function is a fundamental part of biochem-
istry. In spite of its importance, we sometimes forget to mention structure -func-
tion relationships, thinking that the concept is obvious from the examples. In this
book we will try and remind you from time to time how the study of structure leads to a
better understanding of function. This is especially important when studying proteins.

In this chapter and the next one we will cover the basic rules of protein structure. In
Chapters 5 and 6, we will learn how enzymes work and how their structure contributes
to the mechanisms of enzyme action.

Before beginning, let’s review the various kinds of proteins. The following list, al-
though not exhaustive, covers most of the important biological functions of proteins:

1. Many proteins function as enzymes, the biochemical catalysts. Enzymes catalyze
nearly all reactions that occur in living organisms.

2. Some proteins bind other molecules for storage and transport. For example, hemo-
globin binds and transports 0 2 and C0 2 in red blood cells and other proteins bind
fatty acids and lipids.

3. Several types of proteins serve as pores and channels in membranes, allowing for
the passage of small, charged molecules.

4. Some proteins, such as tubulin, actin, and collagen, provide support and shape to
cells and hence to tissues and organisms.

5. Assemblies of proteins can do mechanical work, such as the movement of flagella,
the separation of chromosomes at mitosis, and the contraction of muscles.

6. Many proteins play a role in information flow in the cell. Some are involved in
translation whereas others play a role in regulating gene expression by binding to
nucleic acids.

7. Some proteins are hormones, which regulate biochemical activities in target cells or
tissues; other proteins serve as receptors for hormones.

"Amino acids are literally raining
down from the sky and if that's
not a big deal then I don't know
what is. "

Max Bernstein,
SETI Institute

KEY CONCEPT

The functions of biochemical molecules
can only be understood by knowing their
structures.

Top: L-Arginine, one of the 20 common amino acids.

56 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

KEY CONCEPT

There are many different kinds of proteins
with many different roles in metabolism
and cell structure.

8. Proteins on the cell surface can act as receptors for various ligands and as modifiers
of cell-cell interactions.

9 . Some proteins have highly specialized functions. For example, antibodies defend
vertebrates against bacterial and viral infections, and toxins, produced by bacteria,
can kill larger organisms.

We begin our study of proteins by exploring the structures and chemical properties
of their constituent amino acids. In this chapter we will also discuss the purification,
analysis, and sequencing of polypeptides.

▲ Spindle fibers. Spindle fibers (green) help
separate chromosomes at mitosis. The fibers
are microtubules formed from the structural
protein tubulin.

h 3 n — ch — coo u

5 2 1

▲ Numbering conventions for amino acids. In

traditional names, the carbon atoms adjacent
to the carboxyl group are identified by the
Greek letters a, /3, y, etc. In the official
IUPAC/IUBMB chemical names or systematic
names, the carbon atom in the carboxyl group
is number 1 and the adjacent carbons are
numbered sequentially. Thus, the a-carbon
atom in traditional names is the carbon 2
atom in systematic names.

The IUPAC-IUBMB website for
Nomenclature and Symbolism for
Amino Acids and Peptides is: www.
chem.qmul.ac.uk/iupac/AminoAcid/.

3.1 General Structure of Amino Acids

All organisms use the same 20 amino acids as building blocks for the assembly of protein
molecules. These 20 amino acids are called the common , or standard , amino acids. De-
spite the limited number of amino acids, an enormous variety of different polypeptides
can be produced by connecting the 20 common amino acids in various combinations.

Amino acids are called amino acids because they are amino derivatives of car-
boxylic acids. In the 20 common amino acids the amino group and the carboxyl group
are bonded to the same carbon atom: the ct-carbon atom. Thus, all of the standard
amino acids found in proteins are a-amino acids. Two other substituents are bound to
the a- carbon — a hydrogen atom and a side chain (R) that is distinctive for each amino
acid. In the chemical names of amino acids, carbon atoms are identified by numbers,
beginning with the carbon atom of the carboxyl group. [The correct chemical name, or
systematic name, follows rules established by the International Union of Pure and Ap-
plied Chemistry (IUPAC) and the International Union of Biochemistry and Molecular
Biology (IUBMB).] If the R group is — CH 3 then the systematic name for that amino
acid would be 2-aminopropanoic acid. (Propanoic acid is CH 3 — CH 2 — COOH.) The
trivial name for CH 3 — CH(NH 2 ) — COOH is alanine. The old nomenclature uses Greek
letters to identify the a-carbon atom and the carbon atoms of the side chain. This
nomenclature identifies the carbon atom relative to the carboxyl group so the carbon
atom of the carboxyl group is not specified, unlike in the systematic nomenclature,
where this carbon atom is number 1 in the numbering system. Biochemists have tradi-
tionally used the old, alternate nomenclature.

Inside a cell, under normal physiological conditions, the amino group is protonated
( — NH 3 ©) because the p K a of this group is close to 9. The carboxyl group is ionized
( — COO®) because the p K a of that group is below 3, as we saw in Section 2.9. Thus, in
the physiological pH range of 6.8 to 7.4, amino acids are zwitterions, or dipolar ions, even
though their net charge may be zero. We will see in Section 3.4 that some side chains can
also ionize. Biochemists always represent the structures of amino acids in the form that is
biologically relevant which is why you will see the zwitterions in the following figures.

Figure 3.1a shows the general three-dimensional structure of an amino acid. Figure
3.1b shows a ball-and-stick model of a representative amino acid, serine, whose side
chain is — CH 2 OH. The first carbon atom that’s directly bound to the carboxylate car-
bon is the a - carbon so the other carbon atoms of a side chain are sequentially labeled /3,
y, 8 , and s, referring to carbons 3, 4, 5, and 6, respectively, in the newer convention. The
systematic name for serine is 2-amino-3-hydroxypropanoic acid.

In 19 of the 20 common amino acids the a-carbon atom is chiral, or asymmetric,
since it has four different groups bonded to it. The exception is glycine, whose R group
is simply a hydrogen atom. The molecule is not chiral because the a-carbon atom is
bonded to two identical hydrogen atoms. The 19 chiral amino acids can therefore exist
as stereoisomers. Stereoisomers are compounds that have the same molecular formula
but differ in the arrangement, or configuration, of their atoms in space. The two
stereoisomers are distinct molecules that can’t be easily converted from one form to the
other since a change in configuration requires the breaking of one or more bonds.
Amino acid stereoisomers are nonsuperimposable mirror images called enantiomers.
Two of the 19 chiral amino acids, isoleucine and threonine, have two chiral carbon
atoms each. Isoleucine and threonine can each form four different stereoisomers.

3.1 General Structure of Amino Acids 57

(a)

H 3 N'

";c

(b)

u-Carboxylate group

a-Carbon

-Side chain

# u-Carbon O Nitrogen

O Carbon O Oxygen

O Hydrogen

a-Amino

group

/3-Carbon

By convention, the mirror-image pairs of amino acids are designated D (for dextro,
from the Latin dexter , “right”) and L (for levo, from the Latin laevus , “left”). The config-
uration of the amino acid in Figure 3.1a is L and that of its mirror image is D. To assign
the stereochemical designation, one draws the amino acid vertically with its a-carboxy-
late group at the top and its side chain at the bottom, both pointing away from the
viewer. In this orientation, the a-amino group of the L isomer is on the left of the a-car-
bon, and that of the D isomer is on the right, as shown in Figure 3.2. (The four atoms at-
tached to the a - carbon occupy the four corners of a tetrahedron much like the bonding
of hydrogen atoms to oxygen in water, as shown in Figure 2.4.)

The 19 chiral amino acids used in the assembly of proteins are all of the L configu-
ration, although a few D-amino acids occur in nature. By convention, amino acids are
assumed to be in the L configuration unless specifically designated D. Often it is conven-
ient to draw the structures of L- amino acids in a form that is stereochemically uncom-
mitted, especially when a correct stereochemical representation is not critical to a given
discussion.

The fact that all living organisms use the same standard amino acids in protein
synthesis is evidence that all species on Earth are descended from a common ancestor.
Like modern organisms, the last common ancestor (LCA) must have used L-amino

(a) (b)

Mirror plane Mirror plane

◄ Figure 3.1

Two representations of an L-amino acid at neu-
tral pH. (a) General structure. An amino acid
has a carboxylate group (whose carbon atom
is designated C-l), an amino group, a hydro-
gen atom, and a side chain (or R group), all
attached to C-2 (the a-carbon). Solid
wedges indicate bonds above the plane of
the paper; dashed wedges indicate bonds
below the plane of the paper. The blunt
ends of wedges are nearer the viewer than
the pointed ends, (b) Ball-and-stick model
of serine (whose R group is ( — CH 2 OH).

▲ Meteorites and amino acids. The Murchi-
son meteorite fell in 1969 near Murchison,
Australia. There are many similar carbona-
ceous meteorites and many of them contain
spontaneously formed amino acids, includ-
ing some of the common amino acids found
in proteins. These amino acids are found in
the meteorites as almost equal mixtures of
the l and d configurations.

L-Serine

D-Serine

0 u-Carbon O Nitrogen

O Carbon O Oxygen

O Hydrogen

H 3 N — C — H

See Section 8.1 for a more complete
description of the convention for
displaying stereoisomers (Fischer
projection).

ch 2 oh

L-Serine

D-Serine

◄ Figure 3.2

Mirror-image pairs of amino acids, (a) Ball-
and-stick models of L-serine and D-serine.
Note that the two molecules are not identi-
cal; they cannot be superimposed, (b) L-Ser-
ine and D-serine. The common amino acids
all have the l configuration.

CHAPTER 3 Amino Acids and the Primary Structures of Proteins

acids and not D-amino acids. Mixtures of L- and D-amino acids are formed under con-
ditions that mimic those present when life first arose on Earth 4 billion years ago and
both enantiomers are found in meteorites and in the vicinity of stars. It is not known
how or why primitive life forms selected L- amino acids from the presumed mixture of
the enantiomers present when life first arose. It’s likely that the first proteins were com-
posed of a small number of simple amino acids and selection of L-amino acids over
D-amino acids was a chance event. Modern living organisms do not select L-amino acids
from a mixture because only the L-amino acids are synthesized in sufficient quantities.
Thus, the predominance of L-amino acids in modern species is due to the evolution of
metabolic pathways that produce L-amino acids and not D-amino acids (Chapter 17).

3.2 Structures of the 20 Common Amino Acids

The structures of the 20 amino acids commonly found in proteins are shown in the fol-
lowing figures as Fischer projections. In Fischer projections, horizontal bonds at a chiral
center extend toward the viewer and vertical bonds extend away (as in Figures 3.1 and 3.2).
Examination of the structures reveals considerable variation in the side chains of the 20
amino acids. Some side chains are nonpolar and thus hydrophobic whereas others are
polar or ionized at neutral pH and are therefore hydrophilic. The properties of the side
Some nonstandard amino acids are chains greatly influence the overall three-dimensional shape, or conformation, of a pro-

described in Section 3.3. tein. F° r example, most of the hydrophobic side chains of a water-soluble protein fold

into the interior giving the protein a compact, globular shape.

Both the three-letter and one-letter abbreviations for each amino acid are shown in
the figures. The three-letter abbreviations are self-evident but the one-letter abbreviations
are less obvious. Several amino acids begin with the same letter so other letters of the
alphabet have to be used in order to provide a unique label; for example, threonine = T,
tyrosine = Y, and tryptophan = W. These labels have to be memorized.

BOX 3.1 FOSSIL DATING BY AMINO ACID RACEMIZATION

Amino acids can spontaneously convert from the D configu-
ration to the L configuration and vice versa. This is a chemical
reaction that usually proceeds through a carbanion interme-
diate.

The racemization reaction is normally very slow but it
can be sped up at high temperatures. For example, the half-
life for conversion of L-aspartate to D-aspartate is about 30
days at 100°C. The half-life of this reaction at 37°C is about
350 years and at 18°C its about 50,000 years.

The amino acid composition of mammalian tooth
enamel can be used to determine the age of a fossil if the av-
erage temperature of the environment is known or can be es-
timated. When the amino acids are first synthesized they are
exclusively of the L configuration. Over time, the amount of
the D enantiomer increases and the d/l ratio can be measured
very precisely.

Fossil dating by measuring amino acid racemization has
been superceded by more reliable methods but it’s an inter-
esting example of a slow chemical reaction. Some organisms
contain specific racemases that catalyze the interconversion
of an L-amino acid and a D-amino acid; for example, bacteria
have alanine racemase for converting L- alanine to D- alanine
(see Section 8.7B). These enzymes catalyze thousands of re-
actions per second.

H 3 N — C — H

° c 0

°,f,o

— > i

h«-c—nh 3 ©

L-Amino acid

Carbanion

D-Amino acid

▲ The Badegoule Jaw from a stone age juvenile. Homo sapiens
(Natural History Museum, Lyon, France)

3.2 Structures of the 20 Common Amino Acids 59

It is important to learn the structures of the standard amino acids because we refer
to them frequently in the chapters on protein structure, enzymes, and protein synthesis.
In the following sections we have grouped the standard amino acids by their general
properties and the chemical structures of their side chains. The side chains fall into the
following chemical classes: aliphatic, aromatic, sulfur-containing, alcohols, positively
charged, negatively charged, and amides. Of the 20 amino acids five are further classi-
fied as highly hydrophobic (blue) and seven are classified as highly hydrophilic (red).
Understanding the classification of the R groups will simplify memorizing the struc-
tures and names.

A. Aliphatic R Groups

Glycine (Gly, G) is the smallest amino acid. Since its R group is simply a hydrogen atom,
the a-carbon of glycine is not chiral. The two hydrogen atoms of the a-carbon of
glycine impart little hydrophobic character to the molecule. We will see that glycine
plays a unique role in the structure of many proteins because its side chain is small
enough to fit into niches that cannot accommodate any other amino acid.

Four amino acids — alanine (Ala, A), valine (Val, V), leucine (Leu, L), and the struc-
tural isomer of leucine, isoleucine (lie, I) — have saturated aliphatic side chains. The side
chain of alanine is a methyl group whereas valine has a three-carbon branched side
chain and leucine and isoleucine each contain a four-carbon branched side chain. Both
the a- and /3-carbon atoms of isoleucine are asymmetric. Because isoleucine has two
chiral centers, it has four possible stereoisomers. The stereoisomer used in proteins
is called L-isoleucine and the amino acid that differs at the /3-carbon is called
L-alloisoleucine (Figure 3.3). The other two stereoisomers are D-isoleucine and
D-alloisoleucine.

Alanine, valine, leucine, and isoleucine play an important role in establishing and
maintaining the three-dimensional structures of proteins because of their tendency to
cluster away from water. Valine, leucine, and isoleucine are known collectively as the
branched chain amino acids because their side chains of carbon atoms contain
branches. All three amino acids are highly hydrophobic and they share biosynthesis and
degradation pathways (Chapter 17).

Proline (Pro, P) differs from the other 19 amino acids because its three-carbon side
chain is bonded to the nitrogen of its a-amino group as well as to the a-carbon creating
a cyclic molecule. As a result, proline contains a secondary rather than a primary amino
group. The heterocyclic pyrrolidine ring of proline restricts the geometry of polypep-
tides sometimes introducing abrupt changes in the direction of the peptide chain. The
cyclic structure of proline makes it much less hydrophobic than valine, leucine, and
isoleucine.

B. Aromatic R Groups

Phenylalanine (Phe, F), tyrosine (Tyr, Y), and tryptophan (Trp, W) have side chains
with aromatic groups. Phenylalanine has a hydrophobic benzyl side chain. Tyrosine is
structurally similar to phenylalanine except that the para hydrogen of phenylalanine is
replaced in tyrosine by a hydroxyl group ( — OH) making tyrosine a phenol. The hy-
droxyl group of tyrosine is ionizable but retains its hydrogen under normal physiological
conditions. The side chain of tryptophan contains a bicyclic indole group. Tyrosine and

coo°

1 ©

H 3 N— C — H

H^c — NH 3

H 3 N»- C-*H

H^c — NH 3

h 3 c — c— h

H — C — CH 3

HjC — C — H

oh 2

CH 2

oh 2

ch 3

L-lsoleucine

D-lsoleucine

L-Alloisoleucine

D-Alloisoleucine

COO

k ©

coo

I©

H 3 N — C — H

H 3 N — C — F

ch 3

Glycine [G]

Alanine [A]

(Gly)

(Ala)

coo 0

COO 0

H 3 N— C — H

H 3 N — C — H

cn 2

h 3 c / \h 3

Valine [V]

Leucine [L]

(Val)

(Leu)

COO 0

H 3 N — C — H

H 3 C — C — H

ch 2

ch 3

Isoleucine [I]
(Me)

Phenylalanine [F]
(Phe)

Tyrosine [Y]
(Tyr)

Tryptophan [W]
(Trp)

coo 0

H,N — C — H

/ \

H 2 C x x CH 2

ch 2

Proline [P]
(Pro)

◄ Figure 3.3

Stereoisomers of isoleucine. Isoleucine and
threonine are the only two common amino
acids with more than one chiral center. The
other DL pair of isoleucine isomers is called
alloleucine. Note that in L-isoleucine the
— NH 3 © and — CH 3 groups are both on the
left in this projection, while in D-isoleucine
they are both on the right, so that
D-isoleucine and L-isoleucine are
mirror images.

CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Wavelength (nm)

▲ UV absorbance of proteins. The peak of ab
sorbance of most proteins peaks at 280 nm.
Most of the absorbance is due to the pres-
ence of tryptophan and tyrosine residues in
the protein.

coo°
© 1

H 3 N — C — H

coo°

H 3 N — C — H

ch 3

Methionine [M]
(Met)

Cysteine [C]
(Cys)

COO®

H,N — C — H

coo 0

H 3 N — C — H

H — C — OH

Serine [S]
(Ser)

ch 3

Threonine [T]
(Thr)

▲ A sulfur bridge. Natural stone bridge,
Puente del Inca, in Mendoza, Argentina.
Over the years the bridge has been covered
with sulfur deposits.

tryptophan are not as hydrophobic as phenylalanine because their side chains include
polar groups (Table 3.1, page 62).

All three aromatic amino acids absorb ultraviolet (UV) light because, unlike
the saturated aliphatic amino acids, the aromatic amino acids contain delocalized
7r-electrons. At neutral pH both tryptophan and tyrosine absorb light at a wavelength
of 280 nm whereas phenylalanine is almost transparent at 280 nm and absorbs light
weakly at 260 nm. Since most proteins contain tryptophan and tyrosine they will absorb
light at 280 nm. Absorbance at 280 nm is routinely used to estimate the concentration
of proteins in solutions.

C. R Groups Containing Sulfur

Methionine (Met, M) and cysteine (Cys, C) are the two amino acids whose side chains
contain a sulfur atom. Methionine contains a nonpolar methyl thioether group in its
side chain and this makes it one of the more hydrophobic amino acids. Methionine
plays a special role in protein synthesis because it is almost always the first amino acid in
a growing polypeptide chain. The structure of cysteine resembles that of alanine with a
hydrogen atom replaced by a sulfhydryl group ( — SH).

Although the side chain of cysteine is somewhat hydrophobic, it is also highly reac-
tive. Because the sulfur atom is polarizable the sulfhydryl group of cysteine can form
weak hydrogen bonds with oxygen and nitrogen. Moreover, the sulfhydryl group of cys-
teine residues in proteins can be a weak acid which allows it to lose its proton to become
a negatively charged thiolate ion. (The p iC a of the sulfhydryl group of the free amino
acid is 8.3 but this can range from 5-10 in proteins.)

A compound called cystine can be isolated when some proteins are hydrolyzed.
Cystine is formed from two oxidized cysteine molecules linked by a disulfide bond
(Figure 3.4). Oxidation of the sulfhydryl groups of cysteine molecules proceeds most
readily at slightly alkaline pH values because the sulfhydryl groups are ionized at high pH.
The two cysteine side chains must be adjacent in three-dimensional space in order to form
a disulfide bond but they don’t have to be close together in the amino acid sequence of the
polypeptide chain. They may even be found in different polypeptide chains. Disulfide
bonds, or disulfide bridges, may stabilize the three-dimensional structures of some pro-
teins by covalently cross-linking cysteine residues in peptide chains. Most proteins do not
contain disulfide bridges because conditions inside the cell do not favor oxidation;
however, many secreted, or extracellular, proteins contain disulfide bridges.

D. Side Chains with Alcohol Groups

Serine (Ser, S) and threonine (Thr, T) have uncharged polar side chains containing
/3-hydroxyl groups. These alcohol groups give a hydrophilic character to the aliphatic

©NH 3

^OOC — CH — CH 2 — SH + HS — CH 2 — CH — COO^
©NH 3

Cysteine Cysteine

Oxidation

Reduction

©nh 3

G OOC — CH — CH 2 — s — s — CH 2 — CH — COO 0 + 2 H®

©NH 3

Cystine

▲ Figure 3.4

Formation of cystine. When oxidation links the sulfhydryl groups of two cysteine molecules, the re-
sulting compound is a disulfide called cystine.

3.2 Structures of the 20 Common Amino Acids 61

BOX 3.2 AN ALTERNATIVE NOMENCLATURE

The RS system of configurational nomenclature is also some-
times used to describe the chiral centers of amino acids. The
RS system is based on the assignment of a priority sequence
to the four groups bound to a chiral carbon atom. Once as-
signed, the group priorities are used to establish the configu-
ration of the molecule. Priorities are numbered 1 through
4 and are assigned to groups according to the following rules:

1. For atoms directly attached to the chiral carbon, the one
with the lowest atomic mass is assigned the lowest prior-
ity (number 4).

2. If there are two identical atoms bound to the chiral car-
bon, the priority is decided by the atomic mass of the
next atoms bound. For example, a — CH 3 group has a
lower priority than a — CH 2 Br group because hydrogen
has a lower atomic mass than bromine.

3. If an atom is bound by a double or triple bond, the atom
is counted once for each formal bond. Thus, — CHO,
with a double-bonded oxygen, has a higher priority than

— CH 2 OH. The order of priority for the most common
groups, from lowest to highest, is — H, — CH 3 ,
— C 6 H 5 , — CH 2 OH, —CHO, — COOH, — COOR,
— NH 2 , — NHR, —OH, —OR, and — SH.

With these rules in mind, imagine the molecule as the
steering wheel of a car, with the group of lowest priority
(numbered 4) pointing away from you (like the steering col-
umn) and the other three groups arrayed around the rim of
the steering wheel. Trace the rim of the wheel, moving from
the group of highest priority to the group of lowest priority
(1, 2, 3). If the movement is clockwise, the configuration is R
(from the Latin rectus , “right-handed”). If the movement is
counterclockwise, the configuration is S (from the Latin,
sinister , “left-handed”). The figure demonstrates the assign-
ment of S configuration to L-serine by the RS system.
l- Cysteine has the opposite configuration, R. The dl system
is used more often in biochemistry because not all amino
acids found in proteins have the same RS designation.

◄ Assignment of configuration by the RS
system, (a) Each group attached to a chiral
carbon is assigned a priority based on atomic
mass, 4 being the lowest priority, (b) By orient-
ing the molecule with the priority 4 group
pointing away (behind the chiral carbon) and
tracing the path from the highest priority group
to the lowest, the absolute configuration can
be established. If the sequence 1, 2, 3 is
clockwise, the configuration is R. If the se-
quence 1, 2, 3 is counterclockwise, the config-
uration is S. L-Serine has the S configuration.

side chains. Unlike the more acidic phenolic side chain of tyrosine the hydroxyl groups
of serine and threonine have the weak ionization properties of primary and secondary
alcohols. The hydroxymethyl group of serine ( — CH 2 OH) does not appreciably ionize
in aqueous solutions; nevertheless, this alcohol can react within the active sites of a
number of enzymes as though it were ionized. Threonine, like isoleucine, has two chiral
centers — the a- and /3-carbon atoms. L-Threonine is the only one of the four stereoiso-
mers that commonly occurs in proteins. (The other stereoisomers are called D-threo-
nine, L-allothreonine, and D-allothreonine.)

E. Positively Charged R Groups

Histidine (His, H), lysine (Lys, K), and arginine (Arg, R) have hydrophilic side chains
that are nitrogenous bases. The side chains can be positively charged at physiological
pH.

The side chain of histidine contains an imidazole ring substituent. The proto-
nated form of this ring is called an imidazolium ion (Section 3.4). At pH 7 most his-
tidines are neutral (base form) as shown in the accompanying figure but the form
with a positively charged side chain is present and it becomes more common at
slightly lower pH.

Lysine is a diamino acid with both a- and e-amino groups. The e-amino group
exists as an alkylammonium ion ( — CH 2 — NH 3 ©) at neutral pH and confers a posi-
tive charge on proteins. Arginine is the most basic of the 20 amino acids because its

coo°

H 3 N — C — H

/n:

coo°

Histidine [H]

H 3 N — C — H

(His)

ch 2

COO®

ch 2

H 3 N — C — H

ch 2

cn 2

©nh 3

ch 2

Lysine [K]

(Lys)

/ c %©

h 2 n nh 2

Arginine [R]
(Arg)

62 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

coo°

H 3 N— c — H

H,N— C — H

ch 2

cn 2

coo^

coo u

Aspartate [D]

Glutamate [E]

(Asp)

(Glu)

COO 0

H 3 N — C — H

/ \

/ %

H 2 N 0

Asparagine [N]

Glutamine [Q]

(Asn)

(Gin)

Table 3.1 Hydropathy scale

Amino acid

Free energy
change of transfer"
(kj mol 1 )

Highly hydrophobic

Isoleucine

3.1

Phenylalanine

2.5

Valine

2.3

Leucine

2.2

Methionine

1.1

Less hydrophobic

Tryptophan

^.S h

Alanine

1.0

Glycine

0.67

Cysteine

0.17

Tyrosine

0.08

Proline

-0.29

Threonine

-0.75

Serine

-1.1

Highly hydrophilic

Histidine

-1.7

Glutamate

-2.6

Asparagine

-2.7

Glutamine

-2.9

Aspartate

-3.0

Lysine

-4.6

Arginine

-7.5

°The free-energy change is for transfer of an
amino acid residue from the interior of a lipid bi-
layer to water.

b On other scales, tryptophan has a lower hy-
dropathy value.

[Adapted from Eisenberg, D., Weiss, R. M., Ter-
williger, T. C., Wilcox, W. (1982). Hydrophobic
moments in protein structure. Faraday Symp.
Chem. Soc. 17:109-120.]

side-chain guanidinium ion is protonated under all conditions normally found within a
cell. Arginine side chains also contribute positive charges in proteins.

F. Negatively Charged R Groups and Their Amide Derivatives

Aspartate (Asp, D) and glutamate (Glu, E) are dicarboxylic amino acids and have nega-
tively charged hydrophilic side chains at pH 7. In addition to a-carboxyl groups, aspar-
tate possesses a /3-carboxyl group and glutamate possesses a y-carboxyl group. Aspar-
tate and glutamate confer negative charges on proteins because their side chains are
ionized at pH 7. Aspartate and glutamate are sometimes called aspartic acid and glu-
tamic acid but under most physiological conditions they are found as the conjugate
bases and, like other carboxylates, have the suffix -ate. Glutamate is probably familiar as
its monosodium salt, monosodium glutamate (MSG), which is used in food as a flavor
enhancer.

Asparagine (Asn, N) and glutamine (Gin, Q) are the amides of aspartic acid and
glutamic acid, respectively. Although the side chains of asparagine and glutamine are
uncharged these amino acids are highly polar and are often found on the surfaces of
proteins where they can interact with water molecules. The polar amide groups of as-
paragine and glutamine can also form hydrogen bonds with atoms in the side chains of
other polar amino acids.

G. The Hydrophobicity of Amino Acid Side Chains

The various side chains of amino acids range from highly hydrophobic, through weakly
polar, to highly hydrophilic. The relative hydrophobicity or hydrophilicity of each
amino acid is called its hydropathy.

There are several ways of measuring hydropathy, but most of them rely on calculat-
ing the tendency of an amino acid to prefer a hydrophobic environment over a hy-
drophilic environment. A commonly used hydropathy scale is shown in Table 3.1.
Amino acids with highly positive hydropathy values are considered hydrophobic
whereas those with the largest negative values are hydrophilic. It is difficult to determine
the hydropathy values of some amino acid residues that lie near the center of the scale.
For example, there is disagreement over the hydropathy of the indole group of trypto-
phan and in some tables tryptophan has a much lower hydropathy value. Conversely,
cysteine can have a higher hydropathy value in some tables.

Hydropathy is an important determinant of protein folding because hydrophobic
side chains tend to be clustered in the interior of a protein and hydrophilic residues
are usually found on the surface (Section 4.10). However, it is not yet possible to pre-
dict accurately whether a given residue will be found in the nonaqueous interior of a
protein or on the solvent-exposed surface. On the other hand, hydropathy measure-
ments of free amino acids can be successfully used to predict which segments of
membrane-spanning proteins are likely to be embedded in a hydrophobic lipid
bilayer (Chapter 9).

3.3 Other Amino Acids and Amino Acid Derivatives

More than 200 different amino acids are found in living organisms. In addition to
the 20 common amino acids covered in the previous section there are three others
that are incorporated into proteins during protein synthesis. The 21st amino acid is
N-formylmethionine which serves as the initial amino acid during protein synthesis in
bacteria (Section 22.5). The 22nd amino acid is selenocysteine which contains selenium
in place of the sulfur of cysteine. It is incorporated into a few proteins in almost every
species. Selenocysteine is formed from serine during protein synthesis. The 23rd amino
acid is pyrrolysine, found in some species of archaebacteria. Pyrrolysine is a modified
form of lysine that is synthesized before being added to a growing polypeptide chain by
the translation machinery.

N-formylmethionine, selenocysteine, and pyrrolysine are incorporated at specific
codons and that’s why they are considered additions to the standard repertoire of pro-
tein precursors. Because of post-translational modifications many complete proteins
have more than the standard 23 amino acids used in protein synthesis (see below).

3.4 Ionization of Amino Acids 63

(a)

u ooc— ch 2 — ch 2 — ch 2 — nh 3

y-Ami nobutyrate
(GABA)

(b) ©

N^NH

Histamine

(c)

i ©

CH — CH 2 — NH 2 — CH 3

Epinephrine

(Adrenaline)

Thyroxine / Triiodothyronine

▲ Figure 3.5

Compounds derived from common amino acids, (a) y-Ami nobutyrate. a derivative of glutamate,
(b) Histamine, a derivative of histidine, (c) Epinephrine, a derivative of tyrosine, (d) Thyroxine
and triiodothyronine, derivatives of tyrosine. Thyroxine contains one more atom of iodine (in
parentheses) than does triiodothyronine.

In addition to the common 23 amino acids that are incorporated into proteins, all
species contain a variety of L-amino acids that are either precursors of the common
amino acids or intermediates in other biochemical pathways. Examples are homocys-
teine, homoserine, ornithine, and citrulline (see Chapter 17). S-Adenosylmethionine
(SAM) is a common methyl donor in many biochemical pathways (Section 7.2). Many
species of bacteria and fungi synthesize D-amino acids that are used in cell walls and in
complex peptide antibiotics such as actinomycin.

Several common amino acids are chemically modified to produce biologically im-
portant amines. These are synthesized by enzyme -catalyzed reactions that include de-
carboxylation and deamination. In the mammalian brain, for example, glutamate is
converted to the neurotransmitter y-aminobutyrate (GABA) (Figure 3.5a). Mammals
can also synthesize histamine (Figure 3.5b) from histidine. Histamine controls the con-
striction of certain blood vessels and also the secretion of hydrochloric acid by the
stomach. In the adrenal medulla, tyrosine is metabolized to epinephrine, also known as
adrenaline (Figure 3.5c). Epinephrine and its precursor, norepinephrine (a compound
whose amino group lacks a methyl substituent), are hormones that help regulate me-
tabolism in mammals. Tyrosine is also the precursor of the thyroid hormones thyroxine
and triiodothyronine (Figure 3.5d). Biosynthesis of the thyroid hormones requires io-
dide. Small amounts of sodium iodide are commonly added to table salt to prevent goi-
ter, a condition of hypothyroidism caused by a lack of iodide in the diet.

Some amino acids are chemically modified after they have been incorporated into
polypeptides. In fact, there are hundreds of known post-translational modifications.
For example, some proline residues in the protein collagen are oxidized to form hydrox-
yproline residues (Section 4.1 1). Another common modification is the addition of com-
plex carbohydrate chains — a process known as glycosylation (Chapters 8 and 22). Many
proteins are phosphorylated, usually by the addition of phosphoryl groups to the side
chains of serine, threonine, or tyrosine (histidine, lysine, cysteine, aspartate, and gluta-
mate can also be phosphorylated). The oxidation of pairs of cysteine residues to form
cystine also occurs after a polypeptide has been synthesized.

3.4 Ionization of Amino Acids

The physical properties of amino acids are influenced by the ionic states of the a-carboxyl
and a-amino groups and of any ionizable groups in the side chains. Each ionizable
group is associated with a specific p K a value that corresponds to the pH at which the

COO

C— N— C— H

A {„,

ch 3

/V-formylmethionine

COO°

H 3 N — C — H

SeH

Selenocysteine

coo 0

H 3 N — C — H

Pyrrolysine

64 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

BOX 3.3 COMMON NAMES OF AMINO ACIDS

Alanine:

probably from aldehyde + “an” (for con-
venience) + amine (1849)

Methionine:

side chain is a sulfur (Greek theion ) atom
with a methyl group (1928)

Arginine:

crystallizes as a silver salt, from Latin

Phenylalanine:

alanine with a phenyl group (1883)

argentum (silver) (1886)

Proline:

a corrupted form of “pyrrolidine” because

Asparagine:

first isolated from asparagus (1813)

it forms a pyrrolidine ring (1904)

Aspartate:

similar to asparagine (1836)

Serine:

from the Latin sericum (silk), serine is com-

Glutamate:

first identified in the plant protein gluten

mon in silk (1865)

(1866)

Threonine:

similar to the four- carbon sugar threose

Glutamine:

similar to glutamate (1866)

(1936)

Glycine:

from the Greek glykys (sweet), tastes sweet
(1848)

Tryptophan:

isolated from a tryptic digest of protein 1
Greek phanein (to appear) (1890)

Cysteine:

from the Greek kystis (bladder), discovered
in bladder stones (1882)

Tyrosine:

found in cheese, from the Greek tyros
(cheese) (1890)

Histidine:

first isolated from sturgeon sperm, named
for the Greek histidin (tissue) (1896)

Valine:

derivative of valeric acid from the plant
genus Valeriana (1906)

Isoleucine:

isomer of leucine

Sources: Oxford English Dictionary 2nd ed., and Leung, S.H. (2000) Amino

Leucine:

Lysine:

from the Greek leukos (white), forms white
crystals (1820)

product of protein hydrolysis, from the
Greek lysis (loosening) (1891)

acids, aromatic compounds, and carboxylic acids: how did they get their
common names? /. Chem. Educ. 77: 48-49.

KEY CONCEPT

For every acid-base pair the p/fa is the pH
at which the concentrations of the two
forms are equal.

concentrations of the protonated and unprotonated forms are equal (Section 2.9).
When the pH of the solution is below the p K a the protonated form predominates and
the amino acid is then a true acid that is capable of donating a proton. When the pH of
the solution is above the p K a of the ionizable group the unprotonated form of that
group predominates and the amino acid exists as the conjugate base, which is a proton
acceptor. Every amino acid has at least two p K a values corresponding to the ionization
of the ct-carboxyl and a-amino groups. In addition, seven of the common amino acids
have ionizable side chains with additional, measurable p K a values. These values differ
among the amino acids. Thus, at a given pH, amino acids frequently have different net
charges. Many of the modified amino acids have additional ionizable groups contribut-
ing to the diversity of charged amino acid side chains in proteins. Phosphoserine and
phosphotyrosine, for example, will be negatively charged.

Knowing the ionic states of amino acid side chains is important for two reasons.
First, the charged state influences protein folding and the three-dimensional structure of
proteins (Section 4.10). Second, an understanding of the ionic properties of amino acids
in the active site of an enzyme helps one understand enzyme mechanisms (Chapter 6).

The pK a values of amino acids are determined from titration curves such as those
we saw in the previous chapter. The titration of alanine is shown in Figure 3.6. Alanine
has two ionizable groups — the a -carboxyl and the protonated a -amino group. As more
base is added to the solution of acid, the titration curve exhibits two pK a values, at pH
2.4 and pH 9.9. Each pK a value is associated with a buffering zone where the pH of the
solution changes relatively little when more base is added.

The pK a of an ionizable group corresponds to a midpoint of its titration curve. It is
the pH at which the concentration of the acid form (proton donor) exactly equals the
concentration of its conjugate base (proton acceptor). In the example shown in Figure 3.6
the concentrations of the positively charged form of alanine and of the zwitterion are
equal at pH 2.4.

CH 3 ch 3

i i

©nh 3 — ch— cooh^^©nh 3 — ch— COO 0 + H©

( 3 . 1 )

3.4 Ionization of Amino Acids 65

CH,

H 2 N — CH — COO
(anion)

H ©

CH,

H 3 N — CH — COO
(zwitterion)

,©

H ©

CH 3

H 3 N — CH — COOH

(cation)

◄ Figure 3.6

Titration curve for alanine. The first p K a value
is 2.4; the second is 9.9. pl A i a represents
the isoelectric point of alanine.

At pH 9.9 the concentration of the zwitterion equals the concentration of the nega-
tively charged form.

CH 3 ch 3

i I

©NH3 — CH — COO© NH 2 — CH — COO© + H©

(3.2)

KEY CONCEPT

The ionic state of a particular amino acid
side chain is determined by its p K a value
and the pH of the local environment.

Note that in the acid-base pair shown in the first equilibrium (Reaction 3.1) the
zwitterion is the conjugate base of the acid form of alanine. In the second acid-base pair
(Reaction 3.2) the zwitterion is the proton donor, or conjugate acid, of the more basic
form that predominates at higher pH.

One can deduce that the net charge on alanine molecules at pH 2.4 averages +0.5
because there are equal amounts of neutral zwitterion (+/-) and cation (+). The net
charge at pH 9.9 averages -0.5. Midway between pH 2.4 and pH 9.9, at pH 6.15, the av-
erage net charge on alanine molecules in solution is zero. For this reason, pH 6.15 is re-
ferred to as the isoelectric point (pi), or isoelectric pH, of alanine. If alanine were placed
in an electric field at a pH below its pi it would carry a net positive charge (in other
words, its cationic form would predominate), and it would therefore migrate toward the
cathode (the negative electrode). At a pH higher than its pi alanine would carry a net
negative charge and would migrate toward the anode (the positive electrode). At its iso-
electric point (pH = 6.15) alanine would not migrate in either direction.

Histidine contains an ionizable side chain. The titration curve for histidine contains
an additional inflection point that corresponds to the p K a of its side chain (Figure 3.7a).

v Figure 3.7

Ionization of histidine, (a) Titration curve for
histidine. The three p K a values are 1.8, 6.0,
and 9.3. pi H ii S represents the isoelectric
point of histidine, (b) Deprotonation of the
imidazolium ring of the side chain of
histidine.

(b)

coo°

H 3 N — C — H

Imidazolium ion
(protonated form)
of histidine side chain

H©

H ©

COO°

H 3 N — C — H

CH 2

<+ n:
hM

Imidazole

(deprotonated form)
of histidine side chain

66 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Table 3.2 p K a values of acidic and basic
constituents of free amino acids
at 25°C

Amino acid

p/fa value

Carboxyl

group

Amino

group

Side

chain

Glycine

2.4

9.8

Alanine

2.4

9.9

Valine

2.3

9.7

Leucine

2.3

9.7

Isoleucine

2.3

9.8

Methionine

2.1

9.3

Proline

2.0

10.6

Phenylalanine

2.2

9.3

Tryptophan

2.5

9.4

Serine

2.2

9.2

Threonine

2.1

9.1

Cysteine

1.9

10.7

8.4

Tyrosine

2.2

9.2

10.5

Asparagine

2.1

8.7

Glutamine

2.2

9.1

Aspartic acid

2.0

9.9

3.9

Glutamic acid

2.1

9.5

4.1

Lysine

2.2

9.1

10.5

Arginine

1.8

9.0

12.5

Histidine

1.8

9.3

6.0

As is the case with alanine, the first p (1.8) represents the ionization of the a-COOH
carboxyl group and the most basic pi^ a value (9.3) represents the ionization of the a-
amino group. The middle p K a (6.0) corresponds to the deprotonation of the imida-
zolium ion of the side chain of histidine (Figure 3.7b). At pH 7.0 the ratio of imidazole
(conjugate base) to imidazolium ion (conjugate acid) is 10:1. Thus, the protonated and
neutral forms of the side chain of histidine are both present in significant concentra-
tions near physiological pH. A given histidine side chain in a protein may be either pro-
tonated or unprotonated depending on its immediate environment within the protein.
In other words, the actual p K a value of the side-chain group may not be the same as its
value for the free amino acid in solution. This property makes the side chain of histidine
ideal for the transfer of protons within the catalytic sites of enzymes. (A famous exam-
ple is described in Section 6.7c.)

The isoelectric point of an amino acid that contains only two ionizable groups (the
a-amino and the a-carboxyl groups) is the arithmetic mean of its two pfC a values (i.e.,
pi = (pKi + pK 2 )/2). However, for an amino acid that contains three ionizable groups,
such as histidine, one must assess the net charge of each ionic species. The isoelectric
point for histidine lies between the pFC a values on either side of the species with no net
charge, that is, midway between 6.0 and 9.3, or 7.65.

As shown in Table 3.2 the p fC a values of the a-carboxyl groups of free amino acids
range from 1.8 to 2.5. These values are lower than those of typical carboxylic acids such
as acetic acid (p K a = 4.8) because the neighboring — NH 3 © group withdraws electrons
from the carboxylic acid group and this favors the loss of a proton from the ct-carboxyl
group. The side chains, or R groups, also influence the piC a value of the a - carboxyl
group which is why different amino acids have different p K a values. (We have just seen
that the values for histidine and alanine are not the same.)

The a-COOH group of an amino acid is a weak acid. We can use the
Henderson-Hasselbalch equation (Section 2.9) to calculate the fraction of the group
that is ionized at any given pH.

pH = p K a + log

[proton acceptor]
[proton donor]

(3.3)

For a typical amino acid whose cr-COOH group has a p K a of 2.0, the ratio of pro-
ton acceptor (carboxylate anion) to proton donor (carboxylic acid) at pH 7.0 can be
calculated using the Henderson-Hasselbalch equation.

7.0 = 2.0 +

[RCOO 0 ]
° 9 [RCOOH]

(3.4)

In this case, the ratio of carboxylate anion to carboxylic acid is 100,000:1. This
means that under the conditions normally found inside a cell the carboxylate anion is
the predominant species.

The a-amino group of a free amino acid can exist as a free amine, — NH 2 (proton ac-
ceptor) or as a protonated amine, — NH 3 © (proton donor). The p fC a values range from
8.7 to 10.7 as shown in Table 3.2. For an amino acid whose a-amino group has a p K a value
of 10.0 the ratio of proton acceptor to proton donor is 1:1000 at pH 7.0. In other words,
under physiological conditions the a - amino group is mostly protonated and positively
charged. These calculations verify our earlier statement that free amino acids exist pre-
dominantly as zwitterions at neutral pH. They also show that it is inappropriate to draw
the structure of an amino acid with both — CO OH and — NH groups since there is no
pH at which a significant number of molecules contain a protonated carboxyl group and
an unprotonated amino group (see Problem 19). Note that the secondary amino group of
proline (p K a = 10.6) is also protonated at neutral pH so proline — despite the bonding of
the side chain to the a -amino group — is also zwitter ionic at pH 7.

The seven standard amino acids with readily ionizable groups in their side chains
are aspartate, glutamate, histidine, cysteine, tyrosine, lysine, and arginine. Ionization of
these groups obeys the same principles as ionization of the ct-carboxyl and a-amino
groups and the Henderson-Hasselbalch equation can be applied to each ionization. The
ionization of the y-carboxyl group of glutamate (p K a = 4.1) is shown in Figure 3.8a.

3.5 Peptide Bonds Link Amino Acids in Proteins 67

(a)

(b)

h 3 n-

coo°

-C-H

p ch 2

?ch 2

cU ^OH

J©

pK a = 4.1

Carboxylic acid
(protonated form)
of glutamate side chain

COO

,©

H 3 N — C — H

0ch 2

y ch 2

Carboxylate ion
(deprotonated form)
of glutamate side chain

coo°

H 2 N — C — H

cn 2

H ©

cn 2

|h 2

-Z — >

p/C a = 12.5

|h 2

l H2

‘ ^

cn 2

i\\

H®

H 2 N ' © ' nh 2

HN^ NH 2

Guanidinium ion

Guanidine group

(protonated form)

(deprotonated form)

of arginine side chain

▲ Figure 3.8

Ionization of amino acid side chains, (a) Ionization of the protonated y-carboxyl group of glutamate.
The negative charge of the carboxylate anion is delocalized, (b) Deprotonation of the guanidinium
group of the side chain of arginine. The positive charge is delocalized.

Note that the y-carboxyl group is further removed from the influence of the a-ammo-
nium ion and behaves as a weak acid with a piC a of 4.1. This makes it similar in strength
to acetic acid (pFC a = 4.8) whereas the ct-carboxyl group is a stronger acid (pFC a = 2.1).
Figure 3.8b shows the deprotonation of the guanidinium group of the side chain of argi-
nine in a strongly basic solution. Charge delocalization stabilizes the guanidinium ion
contributing to its high p iC a value of 12.5.

As mentioned earlier, the pFC a values of ionizable side chains in proteins can differ
from those of the free amino acids. Two factors cause this perturbation of ionization
constants. First, a-amino and a-carboxyl groups lose their charges once they are linked
by peptide bonds in proteins — consequently, they exert weaker inductive effects on
their neighboring side chains. Second, the position of an ionizable side chain within the
three dimensional structure of a protein can affect its p K a . For example, the enzyme
ribonuclease A has four histidine residues but the side chain of each residue has
a slightly different p K a as a result of differences in their immediate surroundings, or
microenvironments.

3.5 Peptide Bonds Link Amino Acids in Proteins

The linear sequence of amino acids in a polypeptide chain is called the primary structure
of a protein. Higher levels of structure are referred to as secondary, tertiary, and quater-
nary. The structure of proteins is covered more thoroughly in the next chapter but it’s
important to understand peptide bonds and primary structure before discussing some
of the remaining topics in this chapter.

The linkage formed between amino acids is an amide bond called a peptide bond
(Figure 3.9). This linkage can be thought of as the product of a simple condensation re-
action between the ct-carboxyl group of one amino acid and the a-amino group of an-
other. A water molecule is lost from the condensing amino acids in the reaction. (Recall
from Section 2.6 that such simple condensation reactions are extremely unfavorable in
aqueous solutions due to the huge excess of water molecules. The actual pathway of
protein synthesis involves reactive intermediates that overcome this limitation.) Unlike
the carboxyl and amino groups of free amino acids in solution the groups involved in
peptide bonds carry no ionic charges.

Linked amino acids in a polypeptide chain are called amino acid residues. The
names of residues are formed by replacing the ending -ine or -ate with -yl. For example,
a glycine residue in a polypeptide is called glycyl and a glutamate residue is called glutamyl.

The structure of peptide bonds is
described in Section 4.3.

Protein synthesis (translation) is
described in Chapter 22.

68 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Figure 3.9 ►

Peptide bond between two amino acids. The

structure of the peptide linkage can be
viewed as the product of a condensation
reaction in which the a-carboxyl group of
one amino acid condenses with the a-amino
group of another amino acid. The result is a
dipeptide in which the amino acids are
linked by a peptide bond. Here, alanine is
condensed with serine to form alanylserine.

CPU

ch 2 oh

H 3 N — CH — COO° + H 3 N — CH — COO'

hUO<

■©

ch 3 o

CH.OH

N -terminus H 3 N — CH — C — N — CH — COO° C- terminus

Peptide bond

NH 3

H — C— CH 2 — COO°

c =o

c=o

ch 3

▲ Figure 3.10

Aspartame (aspartylphenylalanine methyl
ester).

In the cases of asparagine, glutamine, and cysteine, -yl replaces the final -e to form as-
paraginyl, glutaminyl, and cysteinyl, respectively. The -yl ending indicates that the
residue is an acyl unit (a structure that lacks the hydroxyl of the carboxyl group). The
dipeptide in Figure 3.9 is called alanylserine because alanine is converted to an acyl unit
but the amino acid serine retains its carboxyl group.

The free amino group and free carboxyl group at the opposite ends of a peptide
chain are called the N- terminus (amino terminus) and the C-terminus (carboxyl termi-
nus), respectively. At neutral pH each terminus carries an ionic charge. By convention,
amino acid residues in a peptide chain are numbered from the N-terminus to the
C-terminus and are usually written from left to right. This convention corresponds to
the direction of protein synthesis (Section 22.6). Synthesis begins with the N-terminal
amino acid — almost always methionine (Section 22.5) — and proceeds sequentially to-
ward the C-terminus by adding one residue at a time.

Both the standard three-letter abbreviations for the amino acids (e.g.,
Gly-Arg-Phe-Ala-Lys) and the one-letter abbreviations (e.g., GRFAK) are used to de-
scribe the sequence of amino acid residues in peptides and polypeptides. It’s important
to know both abbreviation systems. The terms dipeptide , tripeptide , oligopeptide , and
polypeptide refer to chains of two, three, several (up to about 20), and many (usually
more than 20) amino acid residues, respectively. A dipeptide contains one peptide
bond, a tripeptide contains two peptide bonds, and so on. As a general rule, each
peptide chain, whatever its length, possesses one free a-amino group and one free
a-carboxyl group. (Exceptions include covalently modified terminal residues and circu-
lar peptide chains.) Note that the formation of a peptide bond eliminates the ioniz-
able a-carboxyl and a-amino groups found in free amino acids. As a result, most of the
ionic charges associated with a protein molecule are contributed by the side chains of
the amino acids. This means that the solubility and ionic properties of a protein are
largely determined by its amino acid composition. Furthermore, the side chains
of the residues interact with each other and these interactions contribute to the three
dimensional shape and stability of a protein molecule (Chapter 4).

Some peptides are important biological compounds and the chemistry of peptides
is an active area of research. Several hormones are peptides; for example, endorphins
are the naturally occurring molecules that modulate pain in vertebrates. Some very sim-
ple peptides are useful as food additives; for example, the sweetening agent aspartame is
the methyl ester of aspartylphenylalanine (Figure 3.10). Aspartame is about 200 times
sweeter than table sugar and is widely used in diet drinks. There are also many peptide
toxins such as those found in snake venom and poisonous mushrooms.

3.6 Protein Purification Techniques

In order to study a particular protein in the laboratory it must be separated from all other
cell components including other, similar proteins. Few analytical techniques will work
with crude mixtures of cellular proteins because they contain hundreds (or thousands) of
different proteins. The purification steps are different for each protein. They are worked

3.6 Protein Purification Techniques 69

out by trying a number of different techniques until a procedure is developed that repro-
ducibly yields highly purified protein that is still biologically active. Purification steps usu-
ally exploit minor differences in the solubilities, net charges, sizes, and binding specificities
of proteins. In this section, we consider some of the common methods of protein purifica-
tion. Most purification techniques are performed at 0°C to 4°C to minimize temperature-
dependent processes such as protein degradation and denaturation (unfolding).

The first step in protein purification is to prepare a solution of proteins. The source
of a protein is often whole cells in which the target protein accounts for less than 0.1%
of the total dry weight. Isolation of an intracellular protein requires that cells be sus-
pended in a buffer solution and homogenized, or disrupted into cell fragments. Under
these conditions most proteins dissolve. (Major exceptions include membrane proteins
which require special purification procedures.) Let’s assume that the desired protein is
one of many proteins in this solution.

One of the first steps in protein purification is often a relatively crude separation
that makes use of the different solubilities of proteins in salt solutions. Ammonium sul-
fate is frequently used in such fractionations. Enough ammonium sulfate is mixed with
the solution of proteins to precipitate the less soluble impurities, which are removed by
centrifugation. The target protein and other more soluble proteins remain in the fluid
called the supernatant fraction. Next, more ammonium sulfate is added to the super-
natant fraction until the desired protein is precipitated. The mixture is centrifuged, the
fluid removed, and the precipitate dissolved in a minimal volume of buffer solution.
Typically, fractionation using ammonium sulfate gives a two- to threefold purification
(i.e., one-half to two-thirds of the unwanted proteins have been removed from the re-
sulting enriched protein fraction). At this point the solvent containing residual ammo-
nium sulfate is exchanged by dialysis for a buffer solution suitable for chromatography.

In dialysis, a protein solution is sealed in a cylinder of cellophane tubing and sus-
pended in a large volume of buffer. The cellophane membrane is semipermeable — high
molecular weight proteins are too large to pass through the pores of the membrane so
proteins remain inside the tubing while low molecular weight solutes (including, in this
case, ammonium and sulfate ions) diffuse out and are replaced by solutes in the buffer.

Column chromatography is often used to separate a mixture of proteins. A cylindrical
column is filled with an insoluble material such as substituted cellulose fibers or syn-
thetic beads. The protein mixture is applied to the column and washed through the ma-
trix of insoluble material by the addition of solvent. As solvent flows through the col-
umn the eluate (the liquid emerging from the bottom of the column) is collected in
many fractions, a few of which are represented in Figure 3.1 la. The rate at which pro-
teins travel through the matrix depends on interactions between matrix and protein.
For a given column different proteins are eluted at different rates. The concentration of
protein in each fraction can be determined by measuring the absorbance of the eluate at
a wavelength of 280 nm (Figure 3.11b). (Recall from Section 3.2B that at neutral pH,
tyrosine and tryptophan absorb UV light at 280 nm.) To locate the target protein the
fractions containing protein must then be assayed, or tested, for biological activity or
some other characteristic property. Column chromatography may be performed under
high pressure using small, tightly packed columns with solvent flow controlled by a
computer. This technique is called HPLC, for high-performance liquid chromatography.

Chromatographic techniques are classified according to the type of matrix. In ion-
exchange chromatography the matrix carries positive charges (anion -exchange resins) or
negative charges (cation -exchange resins). Anion- exchange matrices bind negatively
charged proteins retaining them in the matrix for subsequent elution. Conversely, cation-
exchange materials bind positively charged proteins. The bound proteins can be serially
eluted by gradually increasing the salt concentration in the solvent. As the salt concentra-
tion is increased it eventually reaches a concentration where the salt ions outcompete pro-
teins in binding to the matrix. At this concentration the protein is released and is collected
in the eluate. Individual bound proteins are eluted at different salt concentrations and this
fractionation makes ion-exchange chromatography a powerful tool in protein purification.

Gel-filtration chromatography separates proteins on the basis of molecular size. The
gel is a matrix of porous beads. Proteins that are smaller than the average pore size

▲ There is only one correct way to write the
sequence of a polypeptide- from N-teminus
to C-terminus.

▲ Green mamba ( Dendroapsis angusticeps).

One of the toxins in the venom of this poi-
sonous snake is a large peptide with the
sequence MICYSHKTPQPSATITCEEKT-
CYKKSVRKL PAVVAGRGCGCPSKEMLVAIH
CCRSDKCNE [Viljoen and Botes (1974).
J.Biol.Chem. 249:366]

70 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Figure 3.1 1 ►

Column chromatography, (a) A mixture of
proteins is added to a column containing a
solid matrix. Solvent then flows into the col-
umn from a reservoir. Washed by solvent,
different proteins (represented by red and
blue bands) travel through the column at
different rates, depending on their interac-
tions with the matrix. Eluate is collected in
a series of fractions, a few of which are
shown, (b) The protein concentration of
each fraction is determined by measuring
the absorbance at 280 nm. The peaks corre-
spond to the elution of the protein bands
shown in (a). The fractions are then tested
for the presence of the target protein.

Fractions collected sequentially

▲ Atypical high-performance liquid chro-
matography (HPLC) system in a research lab
(left). The large instrument on the right is a
mass spectrometer (Istituto di Ricerche
Farmacologiche, Milan, Italy)

penetrate much of the internal volume of the beads and are therefore retarded by the
matrix as the buffer solution flows through the column. The smaller the protein, the
later it elutes from the column. Fewer of the pores are accessible to larger protein mole-
cules. Consequently, the largest proteins flow past the beads and elute first.

Affinity chromatography is the most selective type of column chromatography. It re-
lies on specific binding interactions between the target protein and some other mole-
cule that is covalently bound to the matrix of the column. The molecule bound to the
matrix may be a substance or a ligand that binds to a protein in vivo , an antibody that
recognizes the target protein, or another protein that is known to interact with the tar-
get protein inside the cell. As a mixture of proteins passes through the column only the
target protein specifically binds to the matrix. The column is then washed with buffer
several times to rid it of nonspecifically bound proteins. Finally, the target protein can
be eluted by washing the column with a solvent containing a high concentration of salt
that disrupts the interaction between the protein and column matrix. In some cases,
bound protein can be selectively released from the affinity column by adding excess lig-
and to the elution buffer. The target protein preferentially binds to the ligand in solu-
tion instead of the lower concentration of ligand that is attached to the insoluble matrix
of the column. This method is most effective when the ligand is a small molecule. Affin-
ity chromatography alone can sometimes purify a protein 1000- to 10,000-fold.

3.7 Analytical Techniques

Electrophoresis separates proteins based on their migration in an electric field. In
polyacrylamide gel electrophoresis (PAGE) protein samples are placed on a highly cross-
linked gel matrix of polyacrylamide and an electric field is applied. The matrix is

3.7 Analytical Techniques 71

buffered to a mildly alkaline pH so that most proteins are anionic and migrate toward
the anode. Typically, several samples are run at once together with a reference sample.
The gel matrix retards the migration of large molecules as they move in the electric
field. Hence, proteins are fractionated on the basis of both charge and mass.

A modification of the standard electrophoresis technique uses the negatively
charged detergent sodium dodecyl sulfate (SDS) to overwhelm the native charge on
proteins so that they are separated on the basis of mass only. SDS-polyacrylamide gel
electrophoresis (SDS-PAGE) is used to assess the purity and to estimate the molecular
weight of a protein. In SDS-PAGE the detergent is added to the polyacrylamide gel as
well as to the protein samples. A reducing agent is also added to the samples to reduce
any disulfide bonds. The dodecyl sulfate anion, which has a long hydrophobic tail
(CH 3 (CH 2 )ii 0 S 03 ( ^ ) , Figure 2.8) binds to hydrophobic side chains of amino acid
residues in the polypeptide chain. SDS binds at a ratio of approximately one molecule
for every two residues of a typical protein. Since larger proteins bind proportionately
more SDS the charge-to-mass ratios of all treated proteins are approximately the same.
All the SDS-protein complexes are highly negatively charged and move toward the
anode as diagrammed in Figure 3.12a. However, their rate of migration through the gel
is inversely proportional to the logarithm of their mass — larger proteins encounter
more resistance and therefore migrate more slowly than smaller proteins. This sieving
effect differs from gel-filtration chromatography because in gel filtration larger mole-
cules are excluded from the pores of the gel and hence travel faster. In SDS-PAGE all
molecules penetrate the pores of the gel so the largest proteins travel most slowly. The
protein bands that result from this differential migration (Figure 3.13) can be visualized
by staining. Molecular weights of unknown proteins can be estimated by comparing
their migration to the migration of reference proteins on the same gel.

Although SDS-PAGE is primarily an analytical tool, it can be adapted for purifying
proteins. Denatured proteins can be recovered from SDS-PAGE by cutting out the
bands of a gel. The protein is then electroeluted by applying an electric current to allow
the protein to migrate into a buffer solution. After concentration and the removal of
salts such protein preparations can be used for structural analysis, preparation of anti-
bodies, or other purposes.

(a)

Myosin

/3-galactosidase
Bovine serum albumin

Ovalbumin

Carbonic anhydrase
Soybean trypsin inhibitor
Lysozyme
Aprotinin

I Aprotinin*

5 -

~\ 1 1 1 1 1

1 2 3 4 5

Distance migrated (cm)

▲ Figure 3.13

Proteins separated on an SDS-polyacrylamide
gel. (a) Stained proteins after separation. The
high molecular weight proteins are at the top
of the gel. (b) Graph showing the relationship
between the molecular weight of a protein
and the distance it migrates in the gel.

◄ Figure 3.12

SDS-PAGE. (a) An electrophoresis apparatus
includes an SDS-polyacrylamide gel between
two glass plates and buffer in the upper and
lower reservoirs. Samples are loaded into the
wells of the gel, and voltage is applied. Be-
cause proteins complexed with SDS are neg-
atively charged, they migrate toward the
anode, (b) The banding pattern of the pro-
teins after electrophoresis can be visualized
by staining. The smallest proteins migrate
fastest, so the proteins of lowest molecular
weight are at the bottom of the gel.

CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Mass spectrometry, as the name implies, is a technique that determines the mass of a
molecule. The most basic type of mass spectrometer measures the time that it takes for
a charged gas phase molecule to travel from the point of injection to a sensitive detector.
This time depends on the charge of a molecule and its mass and the result is reported as
the mass/charge ratio. The technique has been used in chemistry for almost 100 years
but its application to proteins was limited because, until recently, it was not possible to
disperse charged protein molecules into a gaseous stream of particles.

This problem was solved in the late 1980s with the development of two new types
of mass spectrometry. In electrospray mass spectrometry the protein solution is pumped
through a metal needle at high voltage to create tiny droplets. The liquid rapidly evapo-
rates in a vacuum and the charged proteins are focused on a detector by a magnetic
field. The second new technique is called matrix-assisted laser desorption ionization
(MALDI). In this method the protein is mixed with a chemical matrix and the mixture is
precipitated on a metal substrate. The matrix is a small organic molecule that absorbs
light at a particular wavelength. A laser pulse at the absorption wavelength imparts en-
ergy to the protein molecules via the matrix. The proteins are instantly released from
the substrate (desorbed) and directed to the detector (Figure 3.14). When time-of- flight
(TOF) is measured, the technique is called MALDI-TOF.

Figure 3.14 ►

MALDI-TOF mass spectrometry, (a) A burst
of light releases proteins from the matrix.

(b) Charged proteins are directed toward the
detector by an electric field, (c) The time of
arrival at the detector depends on the mass
and the charge of the protein.

(a)

Metal -
support

o°

o°<f

°Oo

° m

On ,■

°j>o

Laser

° o

Oi 0°

Proteins

Matrix

molecules

3.8 Amino Acid Composition of Proteins 73

The raw data from a mass spectrometry experiment can be quite simple as shown
in Figure 3.14. There, a single species with one positive charge is detected so the
mass/charge ratio gives the mass directly. In other cases the spectra can be more com-
plicated, especially in electrospray mass spectrometry. Often there are several different
charged species and the correct mass has to be calculated by analyzing a collection of
molecules with charges of +1, +2, +3, etc. The spectrum can be daunting when the
source is a mixture of different proteins. Fortunately, there are sophisticated computer
programs that can analyze the data and calculate the correct masses. The current popu-
larity of mass spectrometry owes as much to the development of this software as it does
to the new hardware and new methods of sample preparation.

Mass spectrometry is very sensitive and highly accurate. Often the mass of a protein
can be obtained from picomole (NT 12 mol) quantities that are isolated from an
SDS-PAGE gel. The correct mass can be determined with an accuracy of less than the
mass of a single proton.

3.8 Amino Acid Composition of Proteins

Once a protein has been isolated its amino acid composition can be determined. First,
the peptide bonds of the protein are cleaved by acid hydrolysis, typically using 6 M HC1
(Figure 3.15). Next, the hydrolyzed mixture, or hydrolysate, is subjected to a chromato-
graphic procedure in which each of the amino acids is separated and quantitated, a
process called amino acid analysis. One method of amino acid analysis involves treat-
ment of the protein hydrolysate with phenylisothiocyanate (PITC) at pH 9.0 to generate
phenylthiocarbamoyl (PTC)-amino acid derivatives (Figure 3.16). The PTC-amino
acid mixture is then subjected to HPLC in a column of fine silica beads to which short
hydrocarbon chains have been attached. The amino acids are separated by the hy-
drophobic properties of their side chains. As each PTC-amino acid derivative is eluted
it is detected and its concentration is determined by measuring the absorbance of the
eluate at 254 nm (the peak absorbance of the PTC moiety). Since different PTC-amino
acid derivatives are eluted at different rates the time at which an amino acid derivative
elutes from the column identifies the amino acid relative to known standards. The
amount of each amino acid in the hydrolysate is proportional to the area under its peak.
With this method, amino acid analysis can be performed on samples as small as 1 pico-
mole of a protein that contains approximately 200 residues.

Despite its usefulness, acid hydrolysis cannot yield a complete amino acid analysis.
Since the side chains of asparagine and glutamine contain amide bonds the acid used to
cleave the peptide bonds of the protein also converts asparagine to aspartic acid and
glutamine to glutamic acid. Other limitations of the acid hydrolysis method include
small losses of serine, threonine, and tyrosine. In addition, the side chain of tryptophan
is almost totally destroyed by acid hydrolysis. There are several ways of overcoming
these limitations. For example, proteins can be hydrolyzed to amino acids by enzymes

John B. Fenn (1917-)

Koichi Tanaka (1959-)

▲ John B. Fenn and Koichi Tanaka were
awarded the Nobel Prize in Chemistry in
2002 “for their development of soft
desorption ionisation methods for mass
spectrometric analyses of biological
macromolecules.”

PITC

COO

H R

Amino

acid

R-, O R 2 O R 3

0 I II I II I

H 3 N — CH — C — N — CH — C — N — CH — COOH

H H

2 H 2 0

6 M HCI

pH = 9.0

COO

H HR

H 3 N— CH — COOH + H 3 N — CH — COOH + H 3 N — CH — COOH
▲ Figure 3.15

Acid-catalyzed hydrolysis of a peptide. Incubation with 6 M HCI at 110°C for 16 to 72 hours releases
the constituent amino acids of a peptide.

PTC-amino acid
▲ Figure 3.16

Amino acid treated with phenylisothiocyanate
(PITC). The a-amino group of an amino acid
reacts with phenylisothiocyanate to give a
phenylthiocarbamoyl-amino acid
(PTC-amino acid).

74 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Figure 3.17 ►

HPLC separation of amino acids. Amino acids
obtained from the enzymatic hydrolysis of a
protein are treated with o-phthalaldehyde
and separated by HPLC.

The frequency of amino acids in pro-
teins is correlated with the number of
codons for each amino acid (Section
22 . 1 )

Table 3.3 Amino acid compositions of
proteins

Amino acid

Frequency in
proteins (%)

Highly hydrophobic

lie (1)

5.2

Val (V)

6.6

Leu (L)

9.0

Phe (F)

3.9

Met (M)

2.4

Less hydrophobic

Ala (A)

8.3

Cly (C)

7.2

Cys (C)

1.7

Trp (W)

1.3

Tyr(Y)

3.2

Pro (P)

5.1

Thr (T)

5.8

Ser (S)

6.9

Highly hydrophilic

Asn (N)

4.4

Gin (Q)

4.0

Acidic

Asp (D)

5.3

Glu (E)

6.2

Basic

His (H)

2.2

Lys (K)

5.7

Arg(R)

5.7

Time (mm:ss)

instead of using acid hydrolysis. The free amino acids are then attached to a chemical
that absorbs light in the ultraviolet and the derivatized amino acids are analyzed by
HPLC (Figure 3.17).

Using various analytical techniques the complete amino acid compositions of
many proteins have been determined. Dramatic differences in composition have been
found, illustrating the tremendous potential for diversity based on different combina-
tions of the 20 amino acids.

The amino acid composition (and sequence) of proteins can also be determined
from the sequence of its gene. In fact, these days it is often much easier to clone and se-
quence DNA than it is to purify and sequence a protein. Table 3.3 shows the average fre-
quency of amino acid residues in more than 1000 different proteins whose sequences
are deposited in protein databases. The most common amino acids are leucine, alanine,
and glycine, followed by serine, valine, and glutamate. Tryptophan, cysteine, and histi-
dine are the least abundant amino acids in typical proteins.

If you know the amino acid composition of a protein you can calculate the molec-
ular weight using the molecular weights of the amino acids in Table 3.4. Be sure to sub-
tract the molecular weight of one water molecule for each peptide bond (Section 3.5). You
can get a rough estimate of the molecular weight of a protein by using the average mo-
lecular weight of a residue (= 110). Thus, a protein of 650 amino acid residues has an
approximate relative molecular mass of 71,500 (M r = 71,500).

3.9 Determining the Sequence of Amino Acid Residues

Amino acid analysis provides information on the composition of a protein but not its
primary structure (sequence of residues). In 1950, Pehr Edman developed a technique
that permits removal and identification of one residue at a time from the N-terminus of
a protein. The Edman degradation procedure involves treating a protein at pH 9.0 with
PITC, also known as the Edman reagent. (Recall that PITC can also be used in the meas-
urement of free amino acids as shown in Figure 3.16.) PITC reacts with the free N-termi-
nus of the chain to form a phenylthiocarbamoyl derivative, or PTC-peptide (Figure 3.18,
on the next page). When the PTC-peptide is treated with an anhydrous acid, such as tri-
fluoroacetic acid the peptide bond of the N-terminal residue is selectively cleaved re-
leasing an anilinothiazolinone derivative of the residue. This derivative can be extracted
with an organic solvent, such as butyl chloride, leaving the remaining peptide in the
aqueous phase. The unstable anilinothiazolinone derivative is then treated with aque-
ous acid which converts it to a stable phenylthiohydantoin derivative of the amino acid
that had been the N-terminal residue (PTH-amino acid). The polypeptide chain in the
aqueous phase, now one residue shorter (residue 2 of the original protein is now the N-
terminus), can be adjusted back to pH 9.0 and treated again with PITC. The entire pro-
cedure can be repeated serially using an automated instrument known as a sequenator.
Each cycle yields a PTH-amino acid that can be identified chromatographically, usually
by HPLC.

3.9 Determining the Sequence of Amino Acid Residues 75

The yield of the Edman degradation procedure under carefully controlled condi-
tions approaches 100% and a few picomoles of sample protein can yield sequences of
30 residues or more before further measurement is obscured by the increasing concen-
tration of unrecovered sample from previous cycles of the procedure. For example,
if the Edman degradation procedure had an efficiency of 98% the cumulative yield at
the 30th cycle would be 0.98 30 , or 0.55. In other words, only about half of the
PTH-amino acids generated in the 30th cycle would be derived from the 30th residue
from the N- terminus.

Rt O

N = C = S + H 2 N — C — C— N

H H

Phenylisothiocyanate ^ Y J

(Edman reagent) N-terminal residue

of polypeptide

pH = 9.0

S Rt O O

II I II II

N — C — N — C — C — N — CH — C — N' wx '

I I I I I I

H H H H R 2 H

Phenylthiocarbamoyl-peptide

Table 3.4 Molecular weights of
amino acids

Amino acid

M r

Ala(A)

Arg(R)

174

Asn(N)

132

Asp(D)

133

Cys(C)

121

Gln(O)

146

Glu(E)

147

Gly(G)

His(H)

155

He(l)

131

Leu(L)

131

Lys(K)

146

Met(M)

149

Phe(F)

165

Pro(P)

115

Ser(S)

105

Thr(T)

119

Trp(W)

204

Tyr(Y)

181

Val(V)

117

f 3 ccooh

O-r

■c^c:
\ /

S -C

R i

Anilinothiazolinone derivative

H 3 N — CH — C —

|\| WA,

r 2 h

Polypeptide chain with
n-1 amino acid residues

Aqueous acid

Phenylthiohydantoin derivative
of extracted N-terminal amino acid

Amino acid identified
chromatographically

Returned to alkaline conditions
for reaction with additional
phenylisothiocyanate in the
next cycle of Edman degradation

◄ Figure 3.18

Edman degradation procedure. The N-terminal
residue of a polypeptide chain reacts with
phenylisothiocyanate to give a phenylthio-
carbamoyl-peptide. Treating this derivative
with trifluoroacetic acid (F 3 CC00H) releases
an anilinothiazolinone derivative of the
N-terminal amino acid residue. The
anilinothiazolinone is extracted and treated
with aqueous acid, which rearranges the
derivative to a stable phenylthiohydantoin
derivative that can then be identified
chromatographically. The remainder of the
polypeptide chain, whose new N-terminal
residue was formerly in the second position,
is subjected to the next cycle of Edman
degradation.

CHAPTER 3 Amino Acids and the Primary Structures of Proteins

t Figure 3.19

Protein cleavage by cyanogen bromide (CNBr).

Cyanogen bromide cleaves polypeptide
chains at the C-terminal side of methionine
residues. The reaction produces a peptidyl
homoserine lactone and generates a new
N-terminus.

3.10 Protein Sequencing Strategies

Most proteins contain too many residues to be completely sequenced by Edman degra-
dation proceeding only from the N-terminus. Therefore, proteases (enzymes that cat-
alyze the hydrolysis of peptide bonds in proteins) or certain chemical reagents are used
to selectively cleave some of the peptide bonds of a protein. The smaller peptides formed
are then isolated and subjected to sequencing by the Edman degradation procedure.

The chemical reagent cyanogen bromide (CNBr) reacts specifically with methionine
residues to produce peptides with C-terminal homoserine lactone residues and new
N-terminal residues (Figure 3.19). Since most proteins contain relatively few methion-
ine residues treatment with CNBr usually produces only a few peptide fragments. For
example, reaction of CNBr with a polypeptide chain containing three internal methion-
ine residues should generate four peptide fragments. Each fragment can then be se-
quenced from its N-terminus.

Many different proteases can be used to generate fragments for protein sequenc-
ing. For example, trypsin specifically catalyzes the hydrolysis of peptide bonds on the
carbonyl side of lysine and arginine residues both of which bear positively charged side
chains (Figure 3.20a). Staphylococcus aureus V8 protease catalyzes the cleavage of pep-
tide bonds on the carbonyl side of negatively charged residues (glutamate and aspar-
tate); under appropriate conditions (50 mM ammonium bicarbonate), it cleaves only
glutamyl bonds. Chymotrypsin, a less specific protease, preferentially catalyzes the hy-
drolysis of peptide bonds on the carbonyl side of uncharged residues with aromatic or
bulky hydrophobic side chains, such as phenylalanine, tyrosine, and tryptophan
(Figure 3.20b).

By judicious application of cyanogen bromide, trypsin, S. aureus V8 protease, and
chymotrypsin to individual samples of a large protein one can generate many peptide
fragments of various sizes. These fragments can then be separated and sequenced by
Edman degradation. In the final stage of sequence determination the amino acid se-
quence of a large polypeptide chain can be deduced by lining up matching sequences of
overlapping peptide fragments as illustrated in Figure 3.20c. When referring to an
amino acid residue whose position in the sequence is known it is customary to follow
the residue abbreviation with its sequence number. For example, the third residue of the
peptide shown in Figure 3.20 is called Ala-3.

The process of generating and sequencing peptide fragments is especially impor-
tant in obtaining information about the sequences of proteins whose N-termini are
blocked. For example, the N-terminal a-amino groups of many bacterial proteins are
formylated and do not react at all when subjected to the Edman degradation procedure.
Peptide fragments with unblocked N-termini can be produced by selective cleavage and
then separated and sequenced so that at least some of the internal sequence of the pro-
tein can be obtained.

For proteins that contain disulfide bonds, the complete covalent structure is not
fully resolved until the positions of the disulfide bonds have been established. The posi-
tions of the disulfide cross-links can be determined by fragmenting the intact protein,
isolating the peptide fragments, and determining which fragments contain cystine
residues. The task of determining the positions of the cross-links becomes quite compli-
cated when the protein contains several disulfide bonds.

H 3 N — Gly— Arg— Phe— Ala— Lys — Met— Trp— Val— COO u

BrCN (+ H 2 0)

H 3 N — Gly— Arg— Phe— Ala — Lys— N — C x + H 3 N — Trp — Val— COO u + H 3 CSCN + + Br e

H 2 C

\ /

h 2 c — o

Peptidyl homoserine lactone

3.10 Protein Sequencing Strategies 77

(a) H 3 N— Gly— Arg— Ala— Ser — Phe— Gly— Asn — Lys — Trp— Glu— Val— COO°

Trypsin

H 3 N— Gly — Arg— COO^ + H 3 N — Ala — Ser — Phe — Gly — Asn — Lys — COCr^ + H 3 N — Trp — Glu — Val — COCr^

(b) H 3 N — Gly — Arg— Ala —Ser— Phe — Gly — Asn— Lys —Trp— Glu — Val— COO°

Chymotrypsin

H 3 N— Gly— Arg — Ala — Ser— Phe— COO u + H 3 N — ly — Asn — Lys — Trp— COO u + H 3 N— Glu— Val— COO u

(c)

Gly— Arg

Ala — Ser — Phe — Gly — Asn — Lys

Trp — Glu — Val

Gly— Arg— Ala— Ser— Phe

Gly — Asn — Lys — Trp

Glu— Val

Deducing the amino acid sequence of a particular protein from the sequence of its
gene (Figure 3.21) overcomes some of the technical limitations of direct analytical tech-
niques. For example, the amount of tryptophan can be determined and aspartate and
asparagine residues can be distinguished because they are encoded by different codons.
However, direct sequencing of proteins is still important since it is the only way of de-
termining whether modified amino acids are present or whether amino acid residues
have been removed after protein synthesis is complete.

Researchers frequently want to identify a particular unknown protein. Let’s say you
have displayed human serum proteins on an SDS gel and you note the presence of a
protein band at 67 KDa. What is that protein? Two recent developments have made the
job of identifying unknown proteins much easier — sensitive mass spectrometry and
genome sequences. Let’s see how they work.

First, you isolate the protein by cutting out the unknown protein band and eluting
the 67 KD protein. The next step is to digest the protein with a protease that cuts at spe-
cific sites. Let’s say you choose trypsin, an enzyme that cleaves the peptide bond follow-
ing arginine (R) or lysine (K) residues. After digestion with trypsin you end up with
several dozen peptide fragments all of which end with arginine or lysine.

Next, you subject the peptide mixture to mass spectrometry choosing a method
such as MALDI-TOF where the precise molecular weights of the peptides can be deter-
mined. The resulting spectrum is shown in Figure 3.22. You now have a “fingerprint” of
the unknown protein corresponding to the molecular weights of all the trypsin diges-
tion products.

In many labs the technique of chemical sequencing using Edman degradation has
been replaced by methods using the mass spectrometer. If you wanted to determine the
sequences of each peptide shown in Figure 3.22 your next step would be to fragment
each peptide into various sized pieces and measure the precise molecular weight of each
fragment in the mass spectrometer.

The data can be used to determine the sequence of the peptide. For example, take
the tryptic peptide of M r = 1226.59 shown in Figure 3.22. One of the large pieces
produced by fragmenting this peptide has a molecular weight of 1079.5. The difference

DNA

Protein

i r

n r

AAG AG T G AAC CTGTC-
^ Lys — Ser — Glu — Pro — Val^

▲ Figure 3.20

Cleavage and sequencing of an oligopeptide.

(a) Trypsin catalyzes cleavage of peptides on
the carbonyl side of the basic residues argi-
nine and lysine, (b) Chymotrypsin catalyzes
cleavage of peptides on the carbonyl side of
uncharged residues with aromatic or bulky
hydrophobic side chains, including pheny-
lalanine, tyrosine, and tryptophan, (c) By
using the Edman degradation procedure to
determine the sequence of each fragment
(highlighted in boxes) and then lining up the
matching sequences of overlapping frag-
ments, one can determine the order of the
fragments and thus deduce the sequence of
the entire oligopeptide.

◄ Figure 3.21

Sequences of DNA and protein. The amino acid
sequence of a protein can be deduced from
the sequence of nucleotides in the correspon-
ding gene. A sequence of three nucleotides
specifies one amino acid. A, C, G, and T rep-
resent the nucleotide residues of DNA.

78 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

1657.74 1853.89

118-130 509-524

M r

▲ Figure 3.22

Tryptic fingerprint of a 67 kDa serum protein. The numbers over each peak are the mass of the
fragment. The number below each mass refer to the residues in Figure 3.23 (Adapted from
Detlevuvkaw, Wikipedia entry on peptide mass fingerprinting)

▲ Frederick Sanger (191 8-) Sanger won the
Nobel Prize in Chemistry in 1958 for his work
on sequencing proteins. He was awarded a
second Nobel Prize in Chemistry in 1980 for
developing methods of sequencing DNA.

corresponds to a Phe (F) residue (1226.6 — 1079.5 = 147.1), meaning that Phe (F) is the
residue at one end of the tryptic peptide. Another large fragment might have a molecu-
lar weight of 1098.5 and the difference (1226.6 — 1098.1) is the exact molecular weight
of a Lys (K) residue. Thus, Lys (K) is the residue at the other end of the peptide. This has
to be the C-terminal end since you know that trypsin cleaves after lysine or arginine
residues. You can get the exact sequence of the peptide by analyzing the masses of all
fragments in this manner. One of them will have a molecular weight of 258.0 and that is
almost certainly the dipeptide Glu-Glu (EE). (The actual analysis is a bit more compli-
cated than this but the principle is the same.)

But it’s often not necessary to do the second mass spectrometry analysis in order to
identify an unknown protein. Since your unkown protein is from a species whose
genome has been sequenced you can simply compare the tryptic fingerprint to the pre-
dicted fingerprints of all the proteins encoded by all the genes in the genome. The data-
base consists of a collection of hypothetical peptides produced by analyzing the amino
acid sequence of each protein including proteins of unknown function that are known
only from their sequence. In most cases your collection of peptide masses from the
unknown protein will match only one protein from one of the genes in the database.

In this case, the match is to human serum albumin, a well known serum protein
(Figure 3.23). The masses of several of the peptides correspond to the predicted masses
of the peptides identified in red in the sequence. Take, for example, the peptide of M r =
1226.59 in the output from the tryptic fingerprint. This is exactly the predicted mass of
the peptide from residues 35-44 (FKDLGEENFK). (Note that the first trypsin cleavage
site follows the arginine residue at position 34 and the second cleavage site is after the
lysine residue at position 44.)

A single match is not sufficient to identify an unknown protein. In the example
shown here there are 21 peptide fragments that match the amino acid sequence of
human serum albumin and this is more than sufficient to uniquely identify the protein.

In 1953, Frederick Sanger was the first scientist to determine the complete sequence
of a protein (insulin). In 1958, he was awarded a Nobel Prize for this work. Twenty- two
years later, Sanger won a second Nobel Prize for pioneering the sequencing of nucleic
acids. Today we know the amino acid sequences of thousands of different proteins.
These sequences not only reveal details of the structure of individual proteins but
also allow researchers to identify families of related proteins and to predict the three-
dimensional structure, and sometimes the function, of newly discovered proteins.

3.1 1 Comparisons of the Primary Structures of Proteins Reveal Evolutionary Relationships 79

MKWVTFISLL

ESAENCDKSL

170

KYLYEIARRH

250

ARLSQRFPKA

FLFSSAYSRG

100

HTLFGDKLCT

180

PYFYAP ELLF

260

EFAEVSKLVT

VFRRDAJ KSE
110

VATLRETYGE

190

FAKRYKAAFT

270

DLTKVHTECC

VAHRFKDLGE

ENFKALVLIA

FAQYLQQCPF

EDHVKLVNEV

120

MADCCAKQEP

130

ERNECFLQHK

200

ECCQAADKAA

280

HGDLLECADD

210

CLLPKLDELR

290

RADLAKYICE

140

DDNPNLPRLV

220

DEGKASSAKQ

300

NQDSISSKLK

TEKAKTCVAD

150

RPEVDV MCTA

230

RLKCASLQKF

310

ECCEKPLLEK

160

FHDNEETFLK

240

GERAFKAWAV

320

SHCIAEVEND

330

EMPADLPSLA

410

FKPLVEEPQN

340

ADFVESKDVC

420

LIKQNCELFE

490

LNQLCVLHEK

570

PKATKEQLKA

500

TPVSDRVTKC

350

KNYAEAKDVF

510

CTESLVNRRP

360

LGMFLYEYAR

430

QLGEYKFQNA

440

LLVRYTKKVP

520

CFSALEVDET

370

RHPDYSVVLL

450

QVSTPTLVEV

530

YVPKEFNAET

380

LRLAKTYETT

460

SRNLGKVGSK

540

FTFHADICTL

580

VMDDFAAFVE

590

KCCKADDKET

600

CFAEEPTMRI

610

RERK

390

LEKCCAAADP

400

HECYAKVFDE

470

CCKHPEAKRM

550

SEKERQIKKQ

480

PCAEDYLSVV

560

TALVELVKHK

▲ Figure 3.23

The sequence of human serum albumin. Red residues highlight predicted tryptic peptides and the
ones identified in the tryptic fingerprint (Figure 3.22) are underlined.

3.11 Comparisons of the Primary Structures of
Proteins Reveal Evolutionary Relationships

In many cases workers have obtained sequences of the same protein from a number of dif-
ferent species. The results show that closely related species contain proteins with very simi-
lar amino acid sequences and that proteins from distantly related species are much less sim-
ilar in sequence. The differences reflect evolutionary change from a common ancestral
protein sequence. As more and more sequences were determined it soon became clear that
one could construct a tree of similarities and this tree closely resembled the phylogenetic
trees constructed from morphological comparisons and the fossil record. The evidence
from molecular data was producing independent confirmation of the history of life.

The first sequence-based trees were published almost 50 years ago. One of the earli-
est examples was the tree for cytochrome c — a single polypeptide chain of approxi-
mately 104 residues. It provides us with an excellent example of evolution at the molec-
ular level. Cytochrome c is found in all aerobic organisms and the protein sequences
from distantly related species, such as mammals and bacteria, are similar enough to
confidently conclude that the proteins are homologous. (Different proteins and genes are
defined as homologues if they have descended from a common ancestor. The evidence
for homology is based on sequence similarity.)

The first step in revealing evolutionary relationships is to align the amino acid se-
quences of proteins from a number of species. Figure 3.24 shows an example of such an
alignment for cytochrome c. The alignment reveals a remarkable conservation of
residues at certain positions. For example, every sequence contains a proline at position
30 and a methionine at position 80. In general, conserved residues contribute to the
structural stability of the protein or are essential for its function.

There is selection against any amino acid substitutions at these invariant posi-
tions. A limited number of substitutions are observed at other sites. In most cases, the
allowed substitutions are amino acid residues with similar properties. For example,
position 20 can be occupied by leucine, isoleucine, or valine — these are all hydropho-
bic residues. Similarly, many sites can be occupied by a number of different polar
residues. Some positions are highly variable — residues at these sites contribute very
little to the structure and function of the protein. The majority of observed amino
acid substitutions in homologous proteins are neutral with respect to natural selection.
The fixation of substitutions at such positions during evolution is due to random ge-
netic drift and the phylogenetic tree represents proteins that have the same fuction
even though they have different amino acid sequences.

The function of cytochrome c is
described in Section 14.7.

KEY CONCEPT

Homology is a conclusion that is based
on evidence such as sequence similarity.
Homologous proteins descend from a
common ancestor. There are degrees of
sequence similarity (e.g., 75% identity),
but homology is an all-or-nothing
conclusion. Something is either
homologous or it isn’t.

CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Figure 3.24 ►

Cytochrome c sequences. The sequences of cytochrome c proteins from various species are aligned
to show their similarities. In some cases, gaps (signified by hyphens) have been introduced to im-
prove the alignment. The gaps represent deletions and insertions in the genes that encode these
proteins. For some species, additional residues at the ends of the sequence have been omitted.
Hydrophobic residues are blue and polar residues are red.

The cytochrome c sequences of humans and chimpanzees are identical. This is a re-
flection of their close evolutionary relationship. The monkey and macaque sequences
are very similar to the human and chimpanzee sequences as expected since all four
species are primates. Similarly, the sequences of the plant cytochrome c molecules re-
semble each other much more than they resemble any of the other sequences.

Figure 3.25 illustrates the similarities between cytochrome c sequences in different
species by depicting them as a tree whose branches are proportional in length to the
number of differences in the amino acid sequences of the protein. Species that are closely
related cluster together on the same branches of the tree because their proteins are very
similar. At great evolutionary distances the number of differences may be very large. For
example, the bacterial sequences differ substantially from the eukaryotic sequences
reflecting divergence from a common ancestor that lived several billion years ago. The
tree clearly reveals the three main kingdoms of eukaryotes — fungi, animals, and plants.
(Protist sequences are not included in this tree in order to make it less complicated.)

Note that every species has changed since divurging from their common ancastor.

Debaryomyces
Candida kloeckeri
krusei

Human,

Zebra, chimpanzee
horse v Macaquej Monkey

RabbitK Penguin

Gray /-Chicken, turkey

kangay^-Puck

xoo/ Pl 9 eon .

^-Snapping turtle

Baker's

yeast

Neurospora
crassa

► Figure 3.25

Phylogenetic tree for cytochrome c. The

length of the branches reflects the number
of differences between the sequences of
many cytochrome c proteins. [Adapted from
Schwartz, R. M., and Dayhoff, M. 0.
(1978). Origins of prokaryotes, eukaryotes,
mitochondria, and chloroplasts. Science
199:395-403.]

Human

GDVEKGKK F

IMKCSQCHTV

EKGGKHKTGP

N HGLFGRKT

50 60

GQAPGYSYTA ANKNKG IWG

EDTLMEYLEN

PKKY1PGTKM

IFVG KKKEE

100

RADLIAYLKK ATNE

Chimpanzee

GDVEKGKK F

IMKCSQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAPGYSYTA

ANKNKG IWG

EDTLMEYLEN

PKKYI PGTKM

IFVG

KKKEE

RADLIAYLKK

ATNE

Spider monkey

GDVFKGKR F

IMKCSQCHTV

EKGGKHKTGP

N HGLFGRKT

GQASG FTYTE

ANKNKG IWG

EDTLMEYLEN

PKKYIPGTKM

IFVG

KKKEE

RADLIAYLKK

ATNE

Macaque

GDVEKGKK F

IMKCSQCHTV

EKGGKHKTGP

N HG GRKT

GQAPGYSYTA

ANKNKGITWG

EDTLMEYLEN

PKKYI PGTKM

IFVG

KKKEE

RAD IAYLKK

ATNE

Cow

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAPG = SYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKGE

RED AYLKK

ATNE

Dog

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAPGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKTGE

RADLIAYLKK

ATKE

Gray whale

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAVGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKGE

RADLIAYLKK

ATNE

Horse

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAPGFTYTD

ANKNKGITWK

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKTE

RE DLIAYLKK

ATNE

Zebra

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HG GRKT

GQAPGFSYTD

ANKNKGITWK

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKTE

RED AYLKK

ATNE

Rabbit

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAVGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKDE

RAD IAYLKK

ATNE

Kangaroo

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

NLHG GRKT

GQAPG =TYTD

ANKNKG IWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKGE

RAD IAYLKK

ATNE

Duck

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAEGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKSE

RADLIAYLKD

ATAK

Turkey

GD EKGKK F

VQKCAQCHTV

EKGGKHKTGP

NLHGLFGRKT

GQAEGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKSE

RVDLIAYLKD

ATSK

Chicken

GD EKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAEGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKSE

RVDLIAYLKD

ATSK

Pigeon

GD EKGKK F

VQKCAQCHTV

EKGGKHKTGP

N HGLFGRKT

GQAEG=SYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKAE

RAD IAYLKQ

ATAK

King penguin

GD EKGKK F

VQKCAQCHTV

EKGGKHKTGP

NLHGIFGRKT

GQAEGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKSE

RAD IAYLKD

ATSK

Snapping turtle

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

NLHG GRKT

GQAEG FSYTE

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKAE

RAD IAYLKD

ATSK

Alligator

GDVEKGKK F

VQKCAQCHTV

EKGGKHKTGP

NLHG GRKT

GQAPG FSYTE

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKPE

RADLIAYLKE

ATSN

Bull frog

GDVEKGKK F

VQKCAQCHTV

EKGGKHKVGP

NLYGL1GRKT

GQAAGFSYTD

ANKNKGITWG

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKGE

RQDLIAYLKS

ACSK

Tuna

GDVAKGKKTF

VQKCAQCHTV

ENGGKHKVGP

NLWG GRKT

GQAEG YSYTD

ANKS KGIVWN

EDTLMEYLEN

PKKYIPGTKM

IFAG

KKKGE

RQDLVAYLKS

ATS

Dogfish

GDVEKGKKVF

VQKCAQCHTV

ENGGKHKTGP

NLSGLFGRKT

GQAQGFSYTD

ANKSKG TWQ

QETLR YLEN

PKKYIPGTKM

IFAG KKKSE

RQDLIAYLKK

TAAS

Starfish

GDVEKGKK F

VQRCAQCHTV

EKAGKHKTGP

NLNG GRKT

GQAAGFSYTD

ANRNKG TWK

NETLF EYI EN

PKKYIPGTKM

VFAG

KKQKE

RQDLIAYLEA

ATK

Fruit fly

GDVEKGKKLF

VQRCAQCHTV

EAGGKHKVGP

NLHGLIGRKT

GQAAGFAYTD

ANKAKGITWN

EDTLF EYLEN

PKKYIPGTKM

IFAG

KKPNE

RGD IAYLKS

ATK

Silkmoth

GNAENGKKIF

VQRCAQCHTV

EAGGKHKVGP

NLHGFYGRKT

GQAPGFSYSN

ANKAKGITWG

DDTLF EYLEN

PKKYIPGTKM

VFAG KKANE

RADLIAYLKE

STK

Pumpkin

GNSKAGEK F

KTKCAQCHTV

DKGAGHKQGP

NLNGLFGRQS

GTTPG YSYSA

ANKNRAVIWE

EKTLY DYLLN

PKKYIPGTKM

VFPG

KKPQD

RADLIAYLKE

ATA

Tomato

GNPKAGEK F

KTKCAQCHTV

EKGAGHKEGP

N NGLFGRQS

GTTAG YSYSA

ANKNMAVNWG

ENTLY DYLLN

PKKYIPGTKM

VFPG

KKPQE

RAD IAYLKE

ATA

Arabidopsis

GDAKKGANLF

KTRCAQCHTL

KAGEGNK GP

ELHGLFGRKT

GSVAGYSYTD

ANKQKG EWK

DDTLF EYI EN

PKKYIPGTKM

A GG

KKPKD

RND ITFLEE

ETK

Mung bean

GNSKSGEK F

KTKCAQCHTV

DKGAGHKQGP

NLNG GRQS

GTTAG YSYST

ANKNMAVIWE

E NTLYDYLLN

PKKYIPGTKM

VFPG

KKPQD

RAD IAYLKE

STA

Wheat

GNPDAGAK

KTKCAQCHTV

DAGAGHKQGP

N HGLFGRQS

GTTAG YSYSA

ANKNRAVEWE

E NTLYDYLLN

PKKYIPGTKM

VFPG

KKPQD

RADLIAYLKK

ATSS

Sunflower

GNPTTGEK F

KTKCAQCHTV

EKGAGHKQGP

N NGLFGRQS

GTTPG YSYSA

GNKNKAVI WE

E NTLYDYLLN

PKKYIPGTKM

VFPG

KKPQE

RADLIAYLKT

STA

Yeast

GSAKKGATLF

KTRCLQCHTV

EKGGPHKVGP

N HG IFGRHS

GQAEG YSYTD

AN KKNVLWD

ENNMSEYLTN

PKKYIPGTKM

A GG

KKEKD

RNDLITYLKK

ACE

Debaryomyces

GSEKKGANLF

KTRCLQCHTV

EKGGPHKVGP

N HGVVGRTS

GQAQGFSYTD

ANKKKGVEWT

EQDLSDY EN

PKKYIPGTKM

AFGG

KKAKD

RNDLITYLVK

ATK

Candida

GSEKKGATLF

KTRCLQCHTV

EKGGPHKVGP

N HGVFGRKS

GLAEGYSYTD

ANKKKGVEWT

EQTMSDYLEN

PKKYIPGTKM

AFGG

LKKPKD

RNDLVTYLKK

ATS

Aspergillus

GDAK-GAKLF

QTRCAQCHTV

EAGGPHKVGP

N HGLFGRKT

GQSEGYAYTD

ANKQAGVTWD

ENT LF S YLEN

PKKF 1 PGTKM

AFGG

LKKGKE

RND ITYLKE

STA

Rhodomicrobium

GDPVKGEQVF

KQ-CK CHQV

GPTAKNGVGP

EQNDVFGQKA

GARPGFNYSD

AMKNSGLTWD

EAT LDKYLEN

PKAVVPGTKM

VFVGLKNPQD

RADVIAYLKQ

LSGK

Nitrobacter

GDVEAGKAAF

NK-CKACHE

GESAKNKVGP

ELDGLDGRHS

GAVEGYAYSP

ANKASG TWD

EAEFKEY KD

PKAKVPGTKM

VFAG

IKKDSE

LDNLWAYVSQ

FDKD

Agrobacterium

GDVAKGEAAF

KR-CSACH A

GEGAKNKVGP

Q NG GRTA

GGDPDYNYSN

AMKKAGLVWT

PQELRDFLSA

PKKK PGNKM

ALAGISKPEE

LDN AYLIF

SASSK

Rhodopila

GDPVEGKHLF

HTICLICHT-

D KGRNKVGP

SLYGVVGRHS

G EPGYNYSE

ANI KSGIVWT

PDVLFKYI E H

PQK PGTKM

GYPG-QPDQK

RADIIAYLET

3.1 1 Comparisons of the Primary Structures of Proteins Reveal Evolutionary Relationships

82 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

Summary

1. Proteins are made from 20 standard amino acids each of which
contains an amino group, a carboxyl group, and a side chain, or
R group. Except for glycine, which has no chiral carbon, all amino
acids in proteins are of the L configuration.

2. The side chains of amino acids can be classified according to their
chemical structures — aliphatic, aromatic, sulfur containing, alco-
hols, bases, acids, and amides. Some amino acids are further clas-
sified as having highly hydrophobic or highly hydrophilic side
chains. The properties of the side chains of amino acids are im-
portant determinants of protein structure and function.

3. Cells contain additional amino acids that are not used in protein
synthesis. Some amino acids can be chemically modified to pro-
duce compounds that act as hormones or neurotransmitters. Some
amino acids are modified after incorporation into polypeptides.

4. At pH 7, the o;-carboxyl group of an amino acid is negatively
charged ( — COO®) and the a-amino group is positively charged
( — NH 3 ®). The charges of ionizable side chains depend on both
the pH and their p K a values.

5. Amino acid residues in proteins are linked by peptide bonds. The
sequence of residues is called the primary structure of the protein.

6. Proteins are purified by methods that take advantage of the differ-
ences in solubility, net charge, size, and binding properties of in-
dividual proteins.

7. Analytical techniques such as SDS-PAGE and mass spectrometry
reveal properties of proteins such as molecular weight.

8. The amino acid composition of a protein can be determined
quantitatively by hydrolyzing the peptide bonds and analyzing the
hydrolysate chromatographically.

9. The sequence of a polypeptide chain can be determined by the
Edman degradation procedure in which the N-terminal residues
are successively cleaved and identified.

10. Proteins with very similar amino acid sequences are homolo-
gous — they descend from a common ancestor.

11. A comparison of sequences from different species reveals evolu-
tionary relationships.

Problems

1. Draw and label the stereochemical structure of L-cysteine. Indi-
cate whether it is R or S by referring to Box 3.2 on page 61.

2. Show that the Fischer projection of the common form of threo-
nine (page 60) corresponds to 2 S, 3R-threonine. Draw and name
the three other isomers of threonine.

3. Histamine dihydrochloride is administered to melanoma (skin
cancer) patients in combination with anticancer drugs because it
makes the cancer cells more receptive to the drugs. Draw the
chemical structure of histamine dihydrochloride.

4. Dried fish treated with salt and nitrite has been found to contain
the mutagen 2-chloro-4-methylthiobutanoic acid (CMBA). From
what amino acid is CMBA derived?

H 3 c — .CH

CH 2

3V “ , ^ n 2\ _ , \

5. For each of the following modified amino acid side chains, iden-
tify the amino acid from which it was derived and the type of
chemical modification that has occurred.

(a) — CH 2 0P0 3 ®

(b) — CH 2 CH1COO 0 2 2

6. The tripeptide glutathione (GSH) (y-Glu-Cys-Gly) serves a pro-
tective function in animals by destroying toxic peroxides that are
generated during aerobic metabolic processes. Draw the chemical
structure of glutathione. Note: The y symbol indicates that the
peptide bond between Glu and Cys is formed between the
y-carboxyl of Glu and the amino group of Cys.

7. Melittin is a 26-residue polypeptide found in bee venom. In its
monomeric form, melittin is thought to insert into lipid-rich
membrane structures. Explain how the amino acid sequence of
melittin accounts for this property.

0 1

H 3 N-Gly-Ile-Gly-Ala-Val-Leu-Lys-Val-Leu-Thr-Gly-Leu

Pro-Ala-Leu-Ile-Ser-Trp-Ile-Lys-Arg-Lys-Arg-Gln-Gln-NH 2

8. Calculate the isoelectric points of (a) arginine and (b) glutamate.

9. Oxytocin is a nonapeptide (a nine-residue peptide) hormone in-
volved in the milk- releasing response in lactating mammals. The
sequence of a synthetic version of oxytocin is shown below. What
is the net charge of this peptide at (a) pH 2.0, (b) pH 8.5, and
(c) pH 10.7? Assume that the ionizable groups have the pK a val-
ues listed in Table 3.2. The disulfide bond is stable at pH 2.0, pH
8.5, and pH 10.7. Note that the C-terminus is amidated.

Cys— Phe— lie — Glu— Asn— Cys — Pro— His — Gly — NH 2

10. Draw the following structures for compounds that would occur
during the Edman degradation procedure: (a) PTC-Leu-Ala,

(b) PTH-Ser, (c) PTH-Pro.

11. Predict the fragments that will be generated from the treatment
of the following peptide with (a) trypsin, (b) chymotrypsin, and

Gly-Ala-Trp-Arg-Asp-Ala-Lys-Glu-Phe-Gly-Gln

Problems 83

12. The titration curve for histidine is shown below. The p K a values
are 1.8 ( — COOH), 6.0 (side chain), and 9.3 ( — NH 3 ®).

(a) Draw the structure of histidine at each stage of ionization.

(b) Identify the points on the titration curve that correspond to
the four ionic species.

(d) Identify the point at which the pH equals the ipK a of the side
chain.

(e) Identify the point that indicates complete titration of the side
chain.

(f ) In what pH ranges would histidine be a good buffer?

13 . You have isolated a decapeptide (a 10-residue peptide) called FP,
which has anticancer activity. Determine the sequence of the pep-
tide from the following information. (Note that amino acids are
separated by commas when their sequence is not known.)

(a) One cycle of Edman degradation of intact FP yields 2 mol of
PTH- aspartate per mole of FP.

(b) Treatment of a solution of FP with 2-mercaptoethanol fol-
lowed by the addition of trypsin yields three peptides with
the composition (Ala, Cys, Phe), (Arg, Asp), and (Asp, Cys,
Gly, Met, Phe). The intact (Ala, Cys, Phe) peptide yields
PTH-cysteine in the first cycle of Edman degradation.

(c) Treatment of 1 mol of FP with carboxypeptidase (which
cleaves the C-terminal residue from peptides) yields 2 mol of
phenylalanine.

(d) Treatment of the intact pentapeptide (Asp, Cys, Gly, Met,
Phe) with CNBr yields two peptides with the composition
(homoserine lactone, Asp) and (Cys, Gly, Phe). The (Cys, Gly,
Phe) peptide yields PTH-glycine in the first cycle of Edman
degradation.

14 . A portion of the amino acid sequences for cytochrome c from the
alligator and bullfrog are given (from Figure 3.24).

Amino acids 31-50

Alligator: NLHGLIGRKT GQAPGFSYTE

Bullfrog: NLYGLIGRKT GQAAGFSYTD

(a) Give an example of a substitution involving similar amino
acids.

(b) Give an example of a more radical substitution.

15 . Several common amino acids are modified to produce biologi-
cally important amines. Serotonin is a biologically important
neurotransmitter synthesized in the brain. Low levels of serotonin
in the brain have been linked to conditions such as depression,
aggression, and hyperactivity. From what amino acid is serotonin
derived? Identify the differences in structure between the amino
acid and serotonin.

16 . The structure of thyrotropin-releasing hormone (TRH) is shown
below. TRH is a peptide hormone originally isolated from the ex-
tracts of hypothalamus.

(a) How many peptide bonds are present in TRH?

(b) From what tripeptide is TRH derived?

CK +h 2 ch 2

ch 2 o o h 2 C +H 2 o

\ /II II \ / //

N— HC c— NH — CH — C N— HC— C

H | \

h 2 c nh 2

HC NH

\ /

N=CH

17 . Chirality plays a major role in the development of new pharma-
ceuticals. People with Parkinsons disease have depleted amounts
of dopamine in their brains. In an effort to increase the amount
of dopamine in patients, they are given the drug L-dopa which is
converted to dopamine in the brain. L-Dopa is marketed in an
enantiomerically pure form, (a) Give the RS designation for
L-dopa. (b) From which amino acid are both L-dopa and dopamine
derived?

co 2

84 CHAPTER 3 Amino Acids and the Primary Structures of Proteins

18. Generations of biochemistry students have encountered a ques-
tion like the one below on their final exam.

Calculate the approximate concentration of the uncharged form
of alanine (see below) in a 0.01 M solution of alanine at (a) pH 2.4
(b) pH 6.15 and (c) pH 9.9.

H 2 N — CH— COOH

Can you answer the question without peeking at the solution?

19. A solution of 0.0 1M alanine is adjusted to pH 2.4 by adding
NaOH. What is the concentration of the zwitterion in this solu-
tion? What would it be if the pH was 4.0?

Selected Readings

General

Creighton, T. E. (1993). Proteins: Structures and
Molecular Principles , 2nd ed. (New York: W. H.
Freeman), pp. 1-48.

Greenstein, J. P., and Winitz, M. (1961). Chemistry
of the Amino Acids (New York: John Wiley 8c
Sons).

Kreil, G. (1997). D-Amino Acids in Animal Pep-
tides. Annu. Rev. Biochem. 66:337-345.

Meister, A. (1965). Biochemistry of the Amino
Acids , 2nd ed. (New York: Academic Press).

Protein Purification and Analysis

Hearn, M. T. W. (1987). General strategies in the
separation of proteins by high-performance liquid
chromatographic methods./. Chromatogr. 418:3-26.

Mann, M., Hendrickson, R.C., and Pandry, A.
(2001) Analysis of Proteins and Proteomes by
Mass Spectrometry. Annu. Rev. Biochem.
70:437-473.

Sherman, L. S., and Goodrich, J. A. (1985). The
historical development of sodium dodecyl
sulphate-polyacrylamide gel electrophoresis.
Chem. Soc. Rev. 14:225-236.

Stellwagen, E. (1990). Gel filtration. Methods Enzy-
mol. 182:317-328.

Amino Acid Analysis and Sequencing

Doolittle, R. F. (1989). Similar amino acid se-
quences revisited. Trends Biochem. Sci.

14:244-245.

Han, K. -K., Belaiche, D., Moreau, O., and Briand,
G. (1985). Current developments in stepwise
Edman degradation of peptides and proteins. Int.

J. Biochem. 17:429-445.

Hunkapiller, M. W., Strickler, J. E., and Wilson, K. J.
(1984). Contemporary methodology for protein
structure determination. Science 226:304-31 1.

Ozols, J. (1990). Amino acid analysis. Methods
Enzymol. 182:587-601.

Sanger, F. (1988). Sequences, sequences, and se-
quences. Annu. Rev. Biochem. 57:1-28.

Proteins: Three-Dimensional
Structure and Function

W e saw in the previous chapter that a protein can be described as a chain of
amino acids joined by peptide bonds in a specific sequence. However,
polypeptide chains are not simply linear but are also folded into compact
shapes that contain coils, zigzags, turns, and loops. Over the last 50 years the three-
dimensional shapes, or conformations, of thousands of proteins have been determined. A
conformation is a spatial arrangement of atoms that depends on the rotation of a bond or
bonds. The conformation of a molecule, such as a protein, can change without breaking
covalent bonds whereas the various configurations of a molecule can be changed only by
breaking and re-forming covalent bonds. (Recall that the L and D forms of amino acids
represent different configurations.) Each protein has an astronomical number of poten-
tial conformations. Since every amino acid residue has a number of possible conforma-
tions and since there are many residues in a protein. Nevertheless, under physiological
conditions most proteins fold into a single stable shape known as its native conforma-
tion. A number of factors constrain rotation around the covalent bonds in a polypep-
tide chain in its native conformation. These include the presence of hydrogen bonds
and other weak interactions between amino acid residues. The biological function of a
protein depends on its native three-dimensional conformation.

A protein may be a single polypeptide chain or it may be composed of several
polypeptide chains bound to each other by weak interactions. As a general rule, each
polypeptide chain is encoded by a single gene although there are some interesting ex-
ceptions to this rule. The size of genes and the polypeptides they encode can vary by
more than an order of magnitude. Some polypeptides contain only 100 amino acid
residues with a relative molecular mass of about 11,000 (M r = 11,000) (Recall that the
average relative molecular mass of an amino acid residue of a protein is 110.) On the
other hand, some very large polypeptide chains contain more than 2000 amino acid
residues (M r = 220,000).

From the intensity of the spots near
the centre , we can infer that the pro-
tein molecules are relatively dense
globular bodies , perhaps joined to-
gether by valency bridges , but in any
event separated by relatively large
spaces which contain water. From the
intensity of the more distant spots , it
can be inferred that the arrangement
of atoms inside the protein molecule is
also of a perfectly definite kind , al-
though without the periodicities char-
acterising the fibrous proteins. The ob-
servations are compatible with oblate
spheroidal molecules of diameters about
25 A. and 35 A., arranged in hexago-
nal screw-axis. ... At this stage , such
ideas are merely speculative , but now
that a crystalline protein has been
made to give X-ray photographs , it is
clear that we have the means of check-
ing them and, by examining the struc-
ture of all crystalline proteins , arriving
at a far more detailed conclusion about
protein structure than previous physi-
cal or chemical methods have been
able to give.

— Dorothy Crowfoot Hodgkin (1 934)

Top: Bighorn sheep. The skin, wool, and horns are composed largely of fibrous proteins.

86 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Classes of proteins are described in
the introduction to Chapter 3, and the
various classes of enzymes are
described in Section 5.1.

The terms globular proteins and fibrous
proteins are rarely used in modern sci-
entific publications. There are many
proteins that don’t fit into either category.

In some species, the size and sequence of every polypeptide can be determined
from the sequence of the genome. There are about 4000 different polypeptides in the
bacterium Escherichia coli with an average size of about 300 amino acid residues
(M r = 33,000). The fruit fly Drosophila melanogaster contains about 14,000 different
polypeptides with an average size about the same as that in bacteria. Humans and other
mammals have about 20,000 different polypeptides. The study of large sets of proteins,
such as the entire complement of proteins produced by a cell, is part of a field of study
called proteomics.

Proteins come in a variety of shapes. Many are water-soluble, compact, roughly
spherical macromolecules whose polypeptide chains are tightly folded. Such proteins —
traditionally called globular proteins — characteristically have a hydrophobic interior and
a hydrophilic surface. They possess indentations or clefts that specifically recognize and
transiently bind other compounds. By selectively binding other molecules these pro-
teins serve as dynamic agents of biological action. Many globular proteins are
enzymes — the biochemical catalysts of cells. About 31% of the polypeptides in E. coli are
classical metabolic enzymes such as those described in the next few chapters. Other pro-
teins include various factors, carrier proteins, and regulatory proteins; 12% of the
known proteins in E. coli fall into these categories.

Polypeptides can also be components of large subcellular or extracellular structures
such as ribosomes, flagella and cilia, muscle, and chromatin. Fibrous proteins are a partic-
ular class of structural proteins that provide mechanical support to cells or organisms.
Fibrous proteins are typically assembled into large cables or threads. Examples of
fibrous proteins are a-keratin, the major component of hair and nails, and collagen, the
major protein component of tendons, skin, bones, and teeth. Other examples of structural
proteins include the protein components of viruses, bacteriophages, spores, and pollen.

► Escherichia coli proteins. Proteins from
E. coli cells are separated by two-dimensional
gel electrophoresis. In the first dimension,
the proteins are separated by a pH gradient
where each protein migrates to its isoelec-
tric point. The second dimension separates
proteins by size on an SDS-polyacrylamide
gel. Each spot corresponds to a single
polypeptide. There are about 4000 different
proteins in E. coli, but some of them are
present in very small quantities and can’t be
seen on this 2-D gel. This figure is from the
Swiss-2D PAGE database. You can visit this
site and click on any one of the spots to find
out more about a particular protein.

4.1 There Are Four Levels of Protein Structure 87

Many proteins are either integral components of membranes or membrane-associated
proteins. Membrane proteins account for at least 16% of the polypeptides in E. coli and
a much higher percentage in eukaryotic cells.

This chapter describes the molecular architecture of proteins. We will explore the
conformation of the peptide bond and see that two simple shapes, the a helix and the
/ 3 sheet, are common structural elements in all classes of proteins. We will describe
higher levels of protein structure and discuss protein folding and stabilization. Finally,
we will examine how protein structure is related to function using collagen, hemoglo-
bin, and antibodies as examples. Above all, we will learn that proteins have properties
beyond those of free amino acids. Chapters 5 and 6 describe the role of proteins as en-
zymes. The structures of membrane proteins are examined in more detail in Chapter 9
and proteins that bind nucleic acids are covered in Chapters 20 to 22.

4.1 There Are Four Levels of Protein Structure

Individual protein molecules have up to four levels of structure (Figure 4.1). As noted in
Chapter 3, primary structure describes the linear sequence of amino acid residues in a
protein. The three-dimensional structure of a protein is described by three additional
levels: secondary structure, tertiary structure, and quaternary structure. The forces re-
sponsible for maintaining, or stabilizing, these three levels are primarily noncovalent.

Secondary structure refers to regularities in local conformations maintained by hy-
drogen bonds between amide hydrogens and carbonyl oxygens of the peptide back-
bone. The major secondary structures are a helices, /3 strands, and turns. Cartoons
showing the structures of folded proteins usually represent ct-helical regions by helices
and (3 strands by broad arrows pointing in the N-terminal to C- terminal direction.

Tertiary structure describes the completely folded and compacted polypeptide chain.
Many folded polypeptides consist of several distinct globular units linked by a short
stretch of amino acid residues as shown in Figure 4.1c. Such units are called domains.
Tertiary structures are stabilized by the interactions of amino acid side chains in non-
neighboring regions of the polypeptide chain. The formation of tertiary structure
brings distant portions of the primary and secondary structures close together.

(a) Primary structure
-Ala-Glu-Val-Thr-Asp-Pro-Gly-

Domain

(b) Secondary structure

a helix

/3 sheet

(d) Quaternary structure

◄ Figure 4.1

Levels of protein structure, (a) The linear
sequence of amino acid residues defines the
primary structure, (b) Secondary structure
consists of regions of regularly repeating
conformations of the peptide chain such as
a helices and /3 sheets, (c) Tertiary structure
describes the shape of the fully folded
polypeptide chain. The example shown has
two domains, (d) Quaternary structure refers
to the arrangement of two or more polypep-
tide chains into a multisubunit molecule.

CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Some proteins possess quaternary structure — the association of two or more
polypeptide chains into a multisubunit, or oligomeric, protein. The polypeptide chains
of an oligomeric protein may be identical or different.

4.2 Methods for Determining Protein Structure

As we saw in Chapter 3, the amino acid sequence of polypeptides (i.e., primary struc-
ture) can be determined directly by sequencing the protein or indirectly by sequencing
the gene. The usual technique for determining the three-dimensional conformation of a
protein is X-ray crystallography. In this technique, a beam of collimated (parallel)
X rays is aimed at a crystal of protein molecules. Electrons in the crystal diffract the
X rays that are then recorded on film or by an electronic detector (Figure 4.2). Mathe-
matical analysis of the diffraction pattern produces an image of the electron clouds sur-
rounding atoms in the crystal. This electron density map reveals the overall shape of the
molecule and the positions of each of the atoms in three-dimensional space. By com-
bining these data with the principles of chemical bonding it is possible to deduce the lo-
cation of all the bonds in a molecule and hence its overall structure. The technique of
X-ray crystallography has developed to the point where it is possible to determine the
structure of a protein without precise knowledge of the amino acid sequence. In prac-
tice, knowledge of the primary structure makes fitting of the electron density map
much easier at the stage where chemical bonds between atoms are determined.

Initially, X-ray crystallography was used to study the simple repeating units of fibrous
proteins and the structures of small biological molecules. Dorothy Crowfoot Hodgkin was
one of the early pioneers in the application of X-ray crystallography to biological mole-
cules. She solved the structure of penicillin in 1947 and developed many of the techniques
used in the study of large proteins. Hodgkin received the Nobel Prize in 1964 for deter-
mining the structure of vitamin B 12 and she later published the structure of insulin.

The chief impediment to determining the three-dimensional structure of an entire
protein was the difficulty of calculating atomic positions from the positions and inten-
sities of diffracted X-ray beams. Not surprisingly, the development of X-ray crystallog-
raphy of macromolecules closely followed the development of computers. By 1962,
John C. Kendrew and Max Perutz had elucidated the structures of the proteins myo-
globin and hemoglobin, respectively, using large and very expensive computers at
Cambridge University in the United Kingdom. Their results provided the first insights
into the nature of the tertiary structures of proteins and earned them a Nobel Prize in
1962. Since then, the structures of many proteins have been revealed by X-ray crystal-
lography. In recent years, there have been significant advances in the technology due to
the availability of inexpensive high-speed computers and improvements in producing
focused beams of X rays. The determination of protein structures is now limited mainly

Figure 4.2 ►

X-ray crystallography, (a) Diagram of X rays
diffracted by a protein crystal, (b) X-ray dif-
fraction pattern of a crystal of adult human
deoxyhemoglobin. The location and intensity
of the spots are used to determine the three-
dimensional structure of the protein.

(a)

Source
of X rays

Beam of
collimated
X rays

(b)

Film

4.2 Methods for Determining Protein Structure 89

◄ Bioinformatics in the 1950s. Bror Strand-
berg (left) and Dick Dickerson (right) carry-
ing computer tapes from the EDSAC II
computer center in Cambridge, UK. The
tapes contain X-ray diffraction data from
crystals of myoglobin.

by the difficulty of preparing crystals of a quality suitable for X-ray diffraction and even
that step is mostly carried out by computer- driven robots.

A protein crystal contains a large number of water molecules and it is often possi-
ble to diffuse small ligands such as substrate or inhibitor molecules into the crystal. In
many cases, the proteins within the crystal retain their ability to bind these ligands and
they often exhibit catalytic activity. The catalytic activity of enzymes in the crystalline
state demonstrates that the proteins crystallize in their in vivo native conformations.
Thus, the protein structures solved by X-ray crystallography are accurate representa-
tions of the structures that exist inside cells.

Once the three-dimensional coordinates of the atoms of a macromolecule have
been determined, they are deposited in a data bank where they are available to other
scientists. Biochemists were among the early pioneers in exploiting the Internet to
share data with researchers around the world — the first public domain databases of
biomolecular structures and sequences were established in the late 1970s. Many of the
images in this text were created using data files from the Protein Data Bank (PDB).

Visit the website for information on how
to view three-dimensional structures
and retrieve data files.

◄ Max Perutz (1914-2002) (left) and John
C. Kendrew (1917-1997) (right). Kendrew
determined the structure of myoglobin and
Perutz determined the structure of hemoglo-
bin. They shared the Nobel Prize in 1962.

90 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

(a)

▲ Figure 4.3

Bovine ( Bos taurus ) ribonuclease A. Ribonu-
clease A is a secreted enzyme that hydrolyzes
RNA during digestion, (a) Space-filling model
showing a bound substrate analog in black,
(b) Cartoon ribbon model of the polypeptide
chain showing secondary structure, (c) View
of the substrate-binding site. The substrate
analog (5'-diphosphoadenine-3'-phosphate)
is depicted as a space-filling model, and the
side chains of amino acid residues are shown
as ball-and-stick models. [PDB 1AFK]

Figure 4.4 ►

Bovine ribonuclease A NMR structure. The

figure combines a set of very similar struc-
tures that satisfy the data on atomic interac-
tions. Only the backbone of the polypeptide
chain is shown. Compare this structure with
that in Figure 4.3b. Note the presence of
disulfide bridges (yellow), which are not
shown in the images derived from the X-ray
crystal structure. [PDB 2AAS].

We will list the PDB filename, or accession number, for every protein structure shown in
this text so that you can view the three-dimensional structure on your own computer.

There are many ways of depicting the three-dimensional structure of proteins.
Space-filling models (Figure 4.3a) depict each atom as a solid sphere. Such images re-
veal the dense, closely packed nature of folded polypeptide chains. Space-filling models
of structures are used to illustrate the overall shape of a protein and the surface exposed
to aqueous solvent. One can easily appreciate that the interior of folded proteins is
nearly impenetrable, even by small molecules such as water.

The structure of a protein can also be depicted as a simplified cartoon that empha-
sizes the backbone of the polypeptide chain (Figure 4.3b). In these models, the amino
acid side chains have been eliminated, making it easier to see how the polypeptide folds
into a three-dimensional shape. Such models have the advantage of allowing us to see
into the interior of the protein, and they also reveal elements of secondary structure such
as a helices and / 3 strands. By comparing the structures of different proteins, it is possible
to recognize common folds and patterns that can t be seen in space-filling models.

The most detailed models are those that emphasize the structures of the amino
acid side chains and the various covalent bonds and weak interactions between atoms
(Figure 4.3c). Such detailed models are especially important in understanding how a
substrate binds in the active site of an enzyme. In Figure 4.3c, the backbone is shown in
the same orientation as in Figure 4.3b.

Another technique for analyzing the macromolecular structure of proteins is nu-
clear magnetic resonance (NMR) spectroscopy. This method permits the study of pro-
teins in solution and therefore does not require the painstaking preparation of crystals.
In NMR spectroscopy, a sample of protein is placed in a magnetic field. Certain atomic
nuclei absorb electromagnetic radiation as the applied magnetic field is varied. Because
absorbance is influenced by neighboring atoms, interactions between atoms that are
close together can be recorded. By combining these results with the amino acid se-
quence and known structural constraints it is possible to calculate a number of struc-
tures that satisfy the observed interactions.

Figure 4.4 depicts the complete set of structures for bovine ribonuclease A — the
same protein whose X-ray crystal structure is shown in Figure 4.3. Note that the possible
structures are very similar and the overall shape of the molecule is easily seen. In some
cases, the set of NMR structures may represent fluctuations, or “breathing,” of the pro-
tein in solution. The similarity of the NMR and X-ray crystal structures indicates that the
protein structures found in crystals accurately represent the structure of the protein in
solution but in some cases the structures do not agree. Often this is due to disordered
regions that do not show up in the X-ray crystal structure (Section 4.7D). On very rare
occasions the protein crystallyzes in a conformation that is not the true native form. The
NMR structure is thought to be more accurate.

In general, the NMR spectra for small proteins such as ribonuclease A can be easily
solved but the spectrum of a large molecule can be extremely complex. For this reason, it
is very difficult to determine the structure of larger proteins but the technique is very
powerful for smaller proteins.

4.3 The Conformation of the Peptide Group 91

4.3 The Conformation of the Peptide Group

Our detailed study of protein structure begins with the structure of the peptide bonds
that link amino acids in a polypeptide chain. The two atoms involved in the peptide
bond, along with their four substituents (the carbonyl oxygen atom, the amide hydro-
gen atom, and the two adjacent a-carbon atoms), constitute the peptide group. X-ray
crystallographic analyses of small peptides reveal that the bond between the carbonyl
carbon and the nitrogen is shorter than typical C — N single bonds but longer than typ-
ical C=N double bonds. In addition, the bond between the carbonyl carbon and the
oxygen is slightly longer than typical C=0 double bonds. These measurements reveal
that peptide bonds have some double-bond properties and can best be represented as a
resonance hybrid (Figure 4.5).

Note that the peptide group is polar. The carbonyl oxygen has a partial negative
charge and can serve as a hydrogen acceptor in hydrogen bonds. The nitrogen has a par-
tial positive charge, and the — NH group can serve as a hydrogen donor in hydrogen
bonds. Electron delocalization and the partial double-bond character of the peptide
bond prevent unrestricted free rotation around the C — N bond. As a result, the atoms
of the peptide group lie in the same plane (Figure 4.6). Rotation is still possible around
each N — C a bond and each C a — C bond in the repeating N — C a — C backbone of
proteins. As we will see, restrictions on free rotation around these two additional bonds
ultimately determine the three-dimensional conformation of a protein.

Because of the double-bond nature of the peptide bond, the conformation of the
peptide group is restricted to one of two possible conformations, either trans or cis
(Figure 4.7). In the trans conformation, the two a-carbons of adjacent amino acid
residues are on opposite sides of the peptide bond and at opposite corners of the rectan-
gle formed by the planar peptide group. In the cis conformation, the two a-carbons are
on the same side of the peptide bond and are closer together. The cis and trans confor-
mations arise during protein synthesis when the peptide bond is formed by joining
amino acids to the growing polypeptide chain. The two conformations are not easily
interconverted by free rotation around the peptide bond once it has formed.

The cis conformation is less favorable than the extended trans conformation be-
cause of steric interference between the side chains attached to the two a-carbon atoms.
Consequently, nearly all peptide groups in proteins are in the trans conformation. Rare
exceptions occur, usually at bonds involving the amide nitrogen of proline. Because of
the unusual ring structure of proline, the cis conformation creates only slightly more
steric interference than the trans conformation.

Remember that even though the atoms of the peptide group lie in a plane, rotation is
still possible about the N — C a and C a — C bonds in the repeating N — C a — C backbone.
This rotation is restricted by steric interference between main-chain and side-chain atoms
of adjacent residues. One of the most important restrictions on free rotation is steric in-
terference between carbonyl oxygens on adjacent amino acid residues in the polypeptide

Trans

(a)

a 2

(b)

— C

1 V/

II V/

-ft I

▲ Figure 4.5

Resonance structure of the peptide bond.

(a) In this resonance form, the peptide bond
is shown as a single C — N bond, (b) In this
resonance form, the peptide bond is shown
as a double bond, (c) The actual structure is
best represented as a hybrid of the two reso-
nance forms in which electrons are delocal-
ized over the carbonyl oxygen, the carbonyl
carbon, and the amide nitrogen. Rotation
around the C — N bond is restricted due to
the double-bond nature of the resonance
hybrid form.

H /R2

JC «2

,N.

'C/

R, H

i, ft

O R 3 H

▲ Figure 4.6

Planar peptide groups in a polypeptide chain.

A peptide group consists of the N — H and
C=0 groups involved in formation of the
peptide bond, as well as the a-carbons on
each side of the peptide bond. Two peptide
groups are highlighted in this diagram.

◄ Figure 4.7

Trans and cis conformations of a peptide group.

Nearly all peptide groups in proteins are in
the trans conformation, which minimizes
steric interference between adjacent side
chains. The arrows indicate the direction
from the N- to the C-terminus.

# u-carbon O Hydrogen Q Oxygen

O Carbonyl carbon O Nitrogen O Side chain

92 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Figure 4.8 ►

Rotation around the N — C a and C a — C bonds
that link peptide groups in a polypeptide chain.

(a) Peptide groups in an extended conforma-
tion. (b) Peptide groups in an unstable confor-
mation caused by steric interference between
carbonyl oxygens of adjacent residues. The
van der Waals radii of the carbonyl oxygen
atoms are shown by the dashed lines. The
rotation angle around the N — C a bond is
called (p (phi), and that around the C a — C
bond is called if/ (psi). The substituents of
the outer a-carbons have been omitted for
clarity.

# u-carbon O Hydrogen

O Carbonyl carbon O Nitrogen

Oxygen
Side chain

chain (Figure 4.8). The presence of bulky side chains also restricts free rotation around
the N — C a and C a — C bonds. Proline is a special case — rotation around the N — C a
bond is constrained because it is part of the pyrrolidine ring structure of proline.

The rotation angle around the N — C a bond of a peptide group is designated cp (phi),
and that around the C a — C bond is designated ip (psi). The peptide bond angle is co
(omega). Because rotation around peptide bonds is hindered by their double-bond char-
acter, most of the conformation of the backbone of a polypeptide can be described by cp
and ip. Each of these angles is defined by the relative positions of four atoms of the back-
bone. Clockwise angles are positive, and counterclockwise angles are negative, with each
having a 180° sweep. Thus, each of the rotation angles can range from —180° to +180°.

The biophysicist G. N. Ramachandran and his colleagues constructed space-filling
models of peptides and made calculations to determine which values of and ip are
sterically permitted in a polypeptide chain. Permissible angles are shown as shaded re-
gions in Ramachandran plots of cp versus ip. Figure 4.9a shows the results of theoretical
calculations — the dark, shaded regions represent permissible angles for most residues,
and the lighter areas cover the cp and ip values for smaller amino acid residues where the

(a)

(b)

▲ Figure 4.9

Ramachandran plot, (a) Solid lines indicate the range of permissible cp and if/ values based on molecular models. Dashed lines give the outer limits for
an alanine residue. Large blue dots correspond to values of cp and if/ that produce recognizable conformations such as the a helix and /3 sheets. The
positions shown for the type II turn are for the second and third residues. The white portions of the plot correspond to values of <p and if/ that were
predicted to occur rarely, (b) Observed cp and if/ values in known structures. Crosses indicate values for typical residues in a single protein. Residues in
an a helix are shown in red, /3-strand residues are blue, and others are green.

4.3 The Conformation of the Peptide Group 93

R groups don’t restrict rotation. Blank areas on a Ramachandran plot are nonpermissi-
ble areas, due largely to steric hindrance. The conformations of several types of ideal
secondary structure fall within the shaded areas, as expected.

Another version of a Ramachandran plot is shown in Figure 4.9b. This plot is based
on the observed cp and i/s angles of hundreds of proteins whose structures are known.
The enclosed inner regions represent angles that are found very frequently, and the
outer enclosed regions represent angles that are less frequent. Typical observed angles
for a helices, /3 sheets, and other structures in a protein are plotted. The most important
difference between the theoretical and observed Ramachandran plots is in the region
around 0 °cp and —90°i/j. This region should not be permitted according to the modeling
studies but there are many examples of residues with these angles. It turns out that
steric clashes are prevented in these regions by allowing a small amount of rotation
around the peptide bond. The peptide group does not have to be exactly planar — a little
bit of wiggle is permitted!

Some bulky amino acid residues have smaller permitted areas. Proline is restricted
to a cp value of about —60° to —77° because its N — C a bond is constrained by inclusion
in the pyrrolidine ring of the side chain. In contrast, glycine is exempt from many steric
restrictions because it lacks a /3-carbon. Thus, glycine residues have greater conforma-
tional freedom than other residues and have cp and i/s values that often fall outside the
shaded regions of the Ramachandran plot.

KEY CONCEPT

The three-dimensional conformation of a
polypeptide backbone is defined by the
cp (phi) and i/j (psi) angles of rotation
around each peptide group.

BOX 4.1 FLOWERING IS CONTROLLED BY CIS/TRANS SWITCHES

Almost all peptide groups adopt the trans conformation since
that is the one favored during protein synthesis. It is much
more stable than the cis conformation (with one exception).
Spontaneous switching to the cis conformation is very rare
and it is almost always accompanied by loss of function since
the structure of the protein is severely affected.

However, the activity of some proteins is actually
regulated by conformation changes due to cis/trans isomer-
ization. The change in peptide group conformation invari-
ably takes place at proline residues because the cis conforma-
tion is almost as stable as the trans conformation. This is the
one exception to the rule.

Specific enzymes, called peptidyl prolyl cis/trans iso-
merases, catalyze the interconversion of cis and trans confor-
mation at proline residues by transiently destabilizing the
resonance hybrid structure of the peptide bond and allowing
rotation. One important class of these enzymes recognizes
Ser-Pro and Thr-Pro bonds whenever the serine and threo-
nine residues are phosphorylated. Phosphorylation of amino
acid residues is an important mechanism of regulation by co-
valent modification (see Section 5.9D). The gene for this type
of peptidyl prolyl cis/trans isomerase is called Pinl and it is
present in all eukaryotes.

In the small flowering plant, Arabidopsis thalianna , Pinl
protein acts on some transcription factors that control the tim-
ing of flowering. When threonine residues are phosphorylated,
the transcription factors are recognized by Pinl and the confor-
mation of the Thr-Pro bond is switched from trans to cis. The
resulting conformational change in the structure of the protein
leads to activation of the transcription factors and transcription
of the genes required for producing flowers. Flowering is con-
siderably delayed when the synthesis of peptidyl prolyl cis/trans
isomerase is inhibited by mutations in the Pinl gene.

In humans the cis/trans isomerase encoded by Pinl plays
a role in regulating gene expression by modifying RNA poly-
merase, transcription factors, and other proteins. Mutations in
this gene have been implicated in several hereditary diseases.
The structure of human peptidyl prolyl cis/trans isomerase is
shown in Figure 4.23e.

a Arabidopsis thalianna, also known as thale cress or mouse-ear
cress, is a relative of mustard. It is a favorite model organism in plant
biology because it is easy to grow in the laboratory.

94 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

▲ Linus Pauling (1901-1994), winner of
the Nobel Prize in Chemistry in 1954 and
the Nobel Peace Prize in 1962.

4.4 The « Helix

The a-helical conformation was proposed in 1950 by Linus Pauling and Robert Corey.
They considered the dimensions of peptide groups, possible steric constraints, and op-
portunities for stabilization by formation of hydrogen bonds. Their model accounted
for the major repeat observed in the structure of the fibrous protein a-keratin. This repeat
of 0.50 to 0.55 nm turned out to be the pitch (the axial distance per turn) of the a helix.
Max Perutz added additional support for the structure when he observed a secondary
repeating unit of 0.15 nm in the X-ray diffraction pattern of a-keratin. The 0.15 nm
repeat corresponds to the rise of the a helix (the distance each residue advances the
helix along its axis). Perutz also showed that the a helix was present in hemoglobin,
confirming that this conformation was present in more complex globular proteins.

In theory, an a helix can be either a right- or a left-handed screw. The a helices
found in proteins are almost always right-handed, as shown in Figure 4.10. In an ideal a
helix, the pitch is 0.54 nm, the rise is 0.15 nm, and the number of amino acid residues
required for one complete turn is 3.6 (i.e., approximately 3 2/3 residues: one carbonyl
group, three N — C a — C units, and one nitrogen). Most a helices are slightly distorted
in proteins but they generally have between 3.5 and 3.7 residues per turn.

Right-handed a helix

Pitch

(advance 0.54 nm
per turn)

Rise (advance per
amino acid residue)

% u-carbon
O Carbonyl carbon
O Hydrogen
O Nitrogen

O Oxygen

Axis

O Side chain

▲ Figure 4.10

a Helix. A region of a-helical secondary structure is shown with the N-terminus at the bottom and the C-terminus at the top of the figure. Each
carbonyl oxygen forms a hydrogen bond with the amide hydrogen of the fourth residue further toward the C-terminus of the polypeptide chain.

The hydrogen bonds are approximately parallel to the long axis of the helix. Note that all the carbonyl groups point toward the C-terminus. In an ideal
a helix, equivalent positions recur every 0.54 nm (the pitch of the helix), each amino acid residue advances the helix by 0.15 nm along the long axis of
the helix (the rise), and there are 3.6 amino acid residues per turn. In a right-handed helix the backbone turns in a clockwise direction when viewed
along the axis from its N-terminus. If you imagine that the right-handed helix is a spiral staircase, you will be turning to the right as you walk down the
staircase.

4.4 The a Helix 95

Within an a helix, each carbonyl oxygen (residue n) of the polypeptide backbone is
hydrogen-bonded to the backbone amide hydrogen of the fourth residue further to-
ward the C-terminus (residue n + 4). (The three amino groups at one end of the helix
and the three carbonyl groups at the other end lack hydrogen-bonding partners within
the helix.) Each hydrogen bond closes a loop containing 13 atoms — the carbonyl oxy-
gen, 1 1 backbone atoms, and the amide hydrogen. Thus, an a helix can also be called a
3.6 13 helix based on its pitch and hydrogen-bonded loop size. The hydrogen bonds
that stabilize the helix are nearly parallel to the long axis of the helix.

The ip and ip angles of each residue in an a helix are similar. They cluster around
a stable region of the Ramachandran plot centered at a cp value of —57° and a ip value of
—47° (Figure 4.9). The similarity of these values is what gives the a helix a regular, re-
peating structure. The intramolecular hydrogen bonds between residues n and n + 4
tend to “lock in” rotation around the N — C a and C a — C bonds restricting the ip and ip
angles to a relatively narrow range.

A single intrahelical hydrogen bond would not provide appreciable structural sta-
bility but the cumulative effect of many hydrogen bonds within an a helix stabilizes this
conformation. Hydrogen bonds between amino acid residues are especially stable in the
hydrophobic interior of a protein where water molecules do not enter and therefore
cannot compete for hydrogen bonding. In an a helix, all the carbonyl groups point to-
ward the C-terminus. The entire helix is a dipole with a positive N-terminus and a neg-
ative C-terminus since each peptide group is polar and all the hydrogen bonds point in
the same direction.

The side chains of the amino acids in an a helix point outward from the cylinder
of the helix and they are not involved in the hydrogen bonds that stabilize the a helix
(Figure 4.11). However, the identity of the side chains affects the stability in other
ways. Because of this, some amino acid residues are found in a-helical conformations
more often than others. For example, alanine has a small, uncharged side chain and
fits well into the ct-helical conformation. Alanine residues are prevalent in the a he-
lices of all classes of proteins. In contrast, tyrosine and asparagine with their bulky
side chains are less common in a helices. Glycine, whose side chain is a single hydro-
gen atom, destabilizes a-helical structures since rotation around its a-carbon is so
unconstrained. For this reason, many a helices begin or end with glycine residues.
Proline is the least common residue in an a helix because its rigid cyclic side chain
disrupts the right-handed helical conformation by occupying space that a neighbor-
ing residue of the helix would otherwise occupy. In addition, because it lacks a hydro-
gen atom on its amide nitrogen, proline cannot fully participate in intrahelical hydrogen
bonding. For these reasons, proline residues are found more often at the ends of a helices
than in the interior.

Proteins vary in their a-helical content. In some proteins most of the residues are in
a helices, whereas other proteins contain very little a-helical structure. The average
content of a helix in the proteins that have been examined is 26%. The length of a
helix in a protein can range from about 4 or 5 residues to more than 40 — the average is
about 12.

Many a helices have hydrophilic amino acids on one face of the helix cylinder and
hydrophobic amino acids on the opposite face. The amphipathic nature of the helix is
easy to see when the amino acid sequence is drawn as a spiral called a helical wheel. The
a helix shown in Figure 4.11 can be drawn as a helical wheel representing the helix
viewed along its axis. Because there are 3.6 residues per turn of the helix, the residues
are plotted every 100° along the spiral (Figure 4.12). Note that the helix is a right-handed
screw and it is terminated by a glycine residue at the C-terminal end. The hydrophilic
residues (asparagine, glutamate, aspartate, and arginine) tend to cluster on one side of
the helical wheel.

Amphipathic helices are often located on the surface of a protein with the hy-
drophilic side chains facing outward (toward the aqueous solvent) and the hydropho-
bic side chains facing inward (toward the hydrophobic interior). For example, the helix
shown in Figures 4.1 1 and 4.12 is on the surface of the water-soluble liver enzyme alco-
hol dehydrogenase with the side chains of the first, fifth, and eighth residues

▲ Figure 4.1 1

View of a right-handed a helix. The blue rib-
bon indicates the shape of the polypeptide
backbone. All the side chains, shown as
bal l-and-stick models, project outward from
the helix axis. This example is from residues
lle-355 (bottom) to Gly-365 (top) of horse
liver alcohol dehydrogenase. Some hydrogen
atoms are not shown. [PDB 1ADF].

▲ A right-handed a helix. This helix was
created by Julian Voss-Andreae. It stands
outside Linus Panling’s childhood home in
Portland, Oregon, United States.

96 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Figure 4.12 ►

a helix in horse liver alcohol dehydrogenase.

Highly hydrophobic residues are blue, less
hydrophobic residues are green, and highly
hydrophilic residues are red. (a) Sequence of
amino acids, (b) Helical wheel diagram.

The known frequencies of various
amino acid residues in a helices are
used to predict the secondary structure
based on the primary sequence alone.

▲ Figure 4.14

Leucine zipper region of yeast
(Saccharomyces cerevisiae). GCN4 protein
bound to DNA. GCN4 is a transcription reg-
ulatory protein that binds to specific DNA
sequences. The DNA-binding region consists
of two amphipathic a helices, one from each
of the two subunits of the protein. The side
chains of leucine residues are shown in
a darker blue than the ribbon. Only the
leucine zipper region of the protein is shown
in the figure. [PDB 1YSA].

(isoleucine, phenylalanine, and leucine, respectively) buried in the protein interior
(Figure 4.13).

There are many examples of two amphipathic a helices that interact to produce an
extended coiled-coil structure where the two a helices wrap around each other with
their hydrophobic faces in contact and their hydrophilic faces exposed to solvent. A
common structure in DNA-binding proteins is called a leucine zipper (Figure 4.14). The
name refers to the fact that two a helices are “zippered” together by the hydrophobic
interactions of leucine residues (and other hydrophobic residues) on one side of an
amphipathic helix. The ends of the helices form the DNA-binding region of the protein.

Some proteins contain a few short regions of a 3 10 helix. Like the a helix, the 3 10
helix is right-handed. The carbonyl oxygen of a 3io helix forms a hydrogen bond with the
amide hydrogen of residue n + 3 (as opposed to residue n + 4 in an a helix) so the 3io helix
has a tighter hydrogen-bonded ring structure than the a helix — 10 atoms rather than
13 — and has fewer residues per turn (3.0) and a longer pitch (0.60 nm) (Figure 4.15).

▲ Figure 4.13

Horse ( Equns ferns) liver alcohol dehydrogenase. The amphipathic a helix is highlighted. The side
chains of highly hydrophobic residues are shown in blue, less hydrophobic residues are green, and
charged residues are shown in red. Note that the side chains of the hydrophobic residues are di-
rected toward the interior of the protein and that the side chains of charged residues are exposed to
the surface. [PDB 1ADF].

4.5 (3 Strands and (3 Sheets 97

The 3 10 helix is slightly less stable than the a helix because of steric hindrances and the
awkward geometry of its hydrogen bonds. When a 3 10 helix occurs, it is usually only a
few residues in length and often is the last turn at the C-terminal end of an a helix.
Because of its different geometry, the ip and ip angles of residues in a 3 10 helix occupy a
different region of the Ramachandran plot than the residues of an a helix (Figure 4.9).

4.5 (3 Strands and (3 Sheets

The other common secondary structure is called p structure, a class that includes
/ 3 strands and (3 sheets, p Strands are portions of the polypeptide chain that are almost
fully extended. Each residue in a /3 strand accounts for about 0.32 to 0.34 nm of the
overall length in contrast to the compact coil of an a helix where each residue corre-
sponds to 0.15 nm of the overall length. When multiple P strands are arranged side-by-
side they form p sheets, a structure originally proposed by Pauling and Corey at the
same time they developed a theoretical model of the a helix.

Proteins rarely contain isolated P strands because the structure by itself is not sig-
nificantly more stable than other conformations. However, /3 sheets are stabilized by hy-
drogen bonds between carbonyl oxygens and amide hydrogens on adjacent p strands.
Thus, in proteins, the regions of p structure are almost always found in sheets.

The hydrogen-bonded P strands can be on separate polypeptide chains or on dif-
ferent segments of the same chain. The P strands in a sheet can be either parallel (run-
ning in the same N- to C-terminal direction) (Figure 4.16a) or antiparallel (running in
opposite N- to C-terminal directions) (Figure 4.16b). When the P strands are antiparallel,
the hydrogen bonds are nearly perpendicular to the extended polypeptide chains. Note
that in the antiparallel p sheet, the carbonyl oxygen and the amide hydrogen atoms of
one residue form hydrogen bonds with the amide hydrogen and carbonyl oxygen of a
single residue in the other strand. In the parallel arrangement, the hydrogen bonds are
not perpendicular to the extended chains and each residue forms hydrogen bonds with
the carbonyl and amide groups of two different residues on the adjacent strand.

Parallel sheets are less stable than antiparallel sheets, possibly because the hydrogen
bonds are distorted in the parallel arrangement. The P sheet is sometimes called a
p pleated sheet since the planar peptide groups meet each other at angles, like the folds
of an accordion. As a result of the bond angles between peptide groups, the amino acid

▲ Figure 4.15

The 3 10 helix. In the 3i 0 helix (left) hydrogen
bonds (pink) form between the amide group
of one residue and the carbonyl oxygen of a
residue three positions away. In an a helix
(right) the carbonyl group bonds to an amino
acid residue four positions away.

v Figure 4.16

p Sheets. Arrows indicate the N- to C-terminal
direction of the peptide chain, (a) Parallel (3
sheet. The hydrogen bonds are evenly spaced
but slanted, (b) Antiparallel (3 sheet. The
hydrogen bonds are essentially perpendicular
to the (3 strands, and the space between
hydrogen -bonded pairs is alternately wide
and narrow.

(a)

(b)

- c x.

ON:'

-C^

-c^

.C.

H Y\

/ \

-Cx.

-C.

/ \

\ H

H Y\

R H

I \ H

H V. I

/ \

i — I ^ c ^ „
^ C N

H O

I H l\

c/ C N

n w

I I

O H

" H I

-x' c \ I /N .

^ I N C \C

Hi I ;|

H R O

O R

H I; I

i — I X C X.-

^ C N I C/
I I H |i

R H O

I I

.->c-

I I

C N

I I

R H

■y

CHAPTER 4 Proteins: Three-Dimensional Structure and Function

▲ Figure 4.17

View of two strands of an antiparallel (3 sheet
from influenza virus A neuraminidase. Only the
side chains of the front (3 strand are shown.
The side chains alternate from one side of
the (3 strand to the other side. Both strands
have a right-handed twist. [PDB 1BJI]

KEY CONCEPT

There are only three different kinds of
common secondary structure: a helix,
p strand, and turns.

▲ U-turns are allowed in proteins.

side chains point alternately above and below the plane of the sheet. A typical /3 sheet
contains from two to as many as 15 individual (3 strands. Each strand has an average of
six amino acid residues.

The (3 strands that make up [3 sheets are often twisted and the sheet is usually dis-
torted and buckled. The three-dimensional view of the (3 sheet of ribonuclease A
(Figure 4.3) shows a more realistic view of (3 sheets than the idealized structures in
Figure 4.16.

A view of two strands of a small (3 sheet is shown in Figure 4.17. The side chains of
the amino acid residues in the front strand alternately project to the left and to the right of
(i.e., above and below) the (3 strand, as described above. Typically, (3 strands twist slightly
in a right-hand direction; that is, they twist clockwise as you look along one strand.

The <p and if/ angles of the bonds in a [3 strand are restricted to a broad range of val-
ues occupying a large, stable region in the upper left-hand corner of the Ramachandran
plot. The typical angles for residues in parallel and antiparallel strands are not identical
(see Figure 4.9). Because most (3 strands are twisted, the <p and if/ angles exhibit a
broader range of values than those seen in the more regular a helix.

Although we usually think of (3 sheets as examples of secondary structure this is
not, strictly speaking, correct. In many cases, the individual (3 strands are located in dif-
ferent regions of the protein and only come together to form the (3 sheet when the pro-
tein adopts its final tertiary conformation. Sometimes the quaternary structure of a
protein gives rise to a large f3 sheet. Some proteins are almost entirely f3 sheets but most
proteins have a much lower (3 - strand content.

In the previous section we noted that amphipathic a helices have hydrophobic
side chains that project outward on one side of the helix. This is the side that interacts
with the rest of the protein creating a series of hydrophobic interactions that help sta-
bilize the tertiary structure. The side chains of / 3 sheets project alternately above and
below the plane of the (3 strands. One surface may consist of hydrophobic side chains
that allow the (3 sheet to lie on top of other hydrophobic residues in the interior of the
protein.

An example of such hydrophobic interactions between two (3 sheets is seen in the
structure of the coat protein of grass pollen grains (Figure 4.18a). This protein is the
major allergen affecting people who are allergic to grass pollen. One surface of each
/ 3 sheet contains hydrophobic side chains and the opposite surface has hydrophilic
side chains. The two hydrophobic surfaces interact to form the hydrophobic core of
the protein and the hydrophilic surfaces are exposed to solvent as shown in Figure
4.18b. This is an example of a (3 sandwich, one of several arrangements of secondary
structural elements that are covered in more detail in the section on tertiary structure
(Section 4.7).

4.6 Loops and Turns

In both an a helix and a (3 strand there are consecutive residues with a similar confor-
mation that is repeated throughout the structure. Proteins also contain stretches of non-
repeating three-dimensional structure. Most of these non-repeating regions of secondary
structure can be characterized as loops or turns since they cause directional changes in the
polypeptide backbone. The conformations of peptide groups in nonrepetitive regions
are constrained just as they are in repetitive regions. They have <p and i[/ values that are
usually well within the permitted regions of the Ramachandran plot and often close
to the values of residues that form a helices or [3 strands.

Foops and turns connect a helices and (3 strands and allow the polypeptide chain
to fold back on itself producing the compact three-dimensional shape seen in the native
structure. As much as one-third of the amino acid residues in a typical protein are
found in such nonrepetitive structures. Loops often contain hydrophilic residues and are
usually found on the surfaces of proteins where they are exposed to solvent and form
hydrogen bonds with water. Some loops consist of many residues of extended nonrepet-
itive structure. About 10% of the residues can be found in such regions.

4.7 Tertiary Structure of Proteins 99

Loops containing only a few (up to five) residues are referred to as turns if they
cause an abrupt change in the direction of a polypeptide chain. The most common
types of tight turns are called reverse turns. They are also called p turns because they
often connect different antiparallel P strands. (Recall that in order to create a P sheet
the polypeptide must fold so that two or more regions of P strand are adjacent to one
another as shown in Figure 4.17.) This terminology is misleading since p turns can also
connect a helices or an a helix and a P strand.

There are two common types of p turn, designated type I and type II. Both types
of turn contain four amino acid residues and are stabilized by hydrogen bonding be-
tween the carbonyl oxygen of the first residue and the amide hydrogen of the fourth
residue (Figure 4.19). Both type I and type II turns produce an abrupt (usually about
180°) change in the direction of the polypeptide chain. In type II turns, the third
residue is glycine about 60% of the time. Proline is often the second residue in both
types of turns.

Proteins contain many turn structures. They all have internal hydrogen bonds that
stabilize the structure and that’s why they can be considered a form of secondary struc-
ture. Turns make up a significant proportion of the structure in many proteins. Some of
the bonds in turn residues have cp and i/j angles that lie outside the “permitted” regions of
a typical Ramachandran plot (Figure 4.9). This is especially true of residues in the third
position of type II turns where there is an abrupt change in the direction of the backbone.
This residue is often glycine so the bond angles can adopt a wider range of values without
causing steric clashes between the side-chain atoms and the backbone atoms. Ramachandran
plots usually show only the permitted regions for all residues except glycine — this is why
the rotation angles of type II turns appear to lie in a restricted area.

(a)

(b)

4.7 Tertiary Structure of Proteins

(b)

>=>

1 )

Tertiary structure results from the folding of a polypeptide (which may already possess
some regions of a helix and P structure) into a closely packed three-dimensional struc-
ture. An important feature of tertiary structure is that amino acid residues that are far
apart in the primary structure are brought together permitting interactions among
their side chains. Whereas secondary structure is stabilized by hydrogen bonding
between amide hydrogens and carbonyl oxygens of the polypeptide backbone, tertiary

▲ Figure 4.18

Structure of PHL P2 from Timothy grass
( Phleum pratense ) pollen, (a) The two short,
two-stranded, antiparallel (3 sheets are high-
lighted in blue and purple to show their ori-
entation within the protein, (b) View of the
/3-sandwich structure in a different orienta-
tion showing hydrophobic residues (blue)
and polar residues (red). A number of
hydrophobic interactions connect the two
(3 sheets. [PDB 1BMW].

(n + 2)

# u-carbon
O p- carbon

O Hydrogen
O Nitrogen

O Oxygen
O Carbon

▲ Figure 4.19

Reverse turns, (a) Type I (3 turn. The structure is stabilized by a hydrogen bond between the carbonyl oxygen of the first N-terminal residue (Phe) and
the amide hydrogen of the fourth residue (Gly). Note the proline residue at position n + 1. (b) Type II (3 turn. This turn is also stabilized by a hydrogen
bond between the carbonyl oxygen of the first N-terminal residue (Val) and the amide hydrogen of the fourth residue (Asn). Note the glycine residue at
position n + 2. [PDB 1AHL (giant sea anemone neurotoxin)].

100

CHAPTER 4 Proteins: Three-Dimensional Structure and Function

structure is stabilized primarily by nonco valent interactions (mostly the hydrophobic
effect) between the side chains of amino acid residues. Disulfide bridges, though cova-
lent, are also elements of tertiary structure they are not part of the primary structure
since they form only after the protein folds.

A. Supersecondary Structures

Supersecondary structures, or motifs, are recognizable combinations of a helices,
/3 strands, and loops that appear in a number of different proteins. Sometimes motifs
are associated with a particular function although structurally similar motifs may have
different functions in different proteins. Some common motifs are shown in Figure 4.20.

One of the simplest motifs is the helix-loop-helix (Figure 4.20a). This structure
occurs in a number of calcium-binding proteins. Glutamate and aspartate residues in
the loop of these proteins form part of the calcium-binding site. In certain DNA-binding
proteins a version of this supersecondary structure is called a helix-turn-helix motif
since the residues that connect the helices form a reverse turn. In these proteins, the
residues of the a helices bind DNA.

The coiled-coil motif consists of two amphipathic a helices that interact through their
hydrophobic edges (Figure 4.20b) as in the leucine zipper example (Figure 4.14). Several
a helices can associate to form a helix bundle (Figure 4.20c). In this case, the individual
a helices have opposite orientations, whereas they are parallel in the coiled-coil motif.

The /3af3 unit consists of two parallel /3 strands linked to an intervening a helix by
two loops (Figure 4.20d). The helix connects the C-terminal end of one (3 strand to the
N-terminal end of the next and often runs parallel to the two strands. A hairpin consists
of two adjacent antiparallel / 3 strands connected by a [3 turn (Figure 4.20e). (One exam-
ple of a hairpin motif is shown in Figure 4.16.)

Figure 4.20 ►

Common motifs. In folded proteins a helices
and strands are commonly connected by
loops and turns to form supersecondary
structures, shown here as two-dimensional
representations. Arrows indicate the N- to
C-terminal direction of the peptide chain.

(a) Helix-loop-helix (b) Coiled coil (c) Helix bundle

(g) Greek key

(h) /3-sandwich

4.7 Tertiary Structure of Proteins

101

The [3 meander motif (Figure 4.20f) is an antiparallel [3 sheet composed of sequen-
tial (3 strands connected by loops or turns. The order of strands in the (3 sheet is the
same as their order in the sequence of the polypeptide chain. The (3 meander sheet may
contain one or more hairpins but, more typically, the strands are joined by larger loops.
The Greek key motif takes its name from a design found on classical Greek pottery. This
is a [3 sheet motif linking four antiparallel (3 strands such that strands 3 and 4 form the
outer edges of the sheet and strands 1 and 2 are in the middle of the sheet. The (3 sandwich
motif is formed when / 3 strands or sheets stack on top of one another (Figure 4.20h). The
figure shows an example of a (3 sandwich where the (3 strands are connected by short
loops and turns, but (3 sandwiches can also be formed by the interaction of two (3 sheets
in different regions of the polypeptide chain, as seen in Figure 4.18.

B. Domains

Many proteins are composed of several discrete, independently folded, compact units
called domains. Domains may consist of combinations of motifs. The size of a domain
varies from as few as 25 to 30 amino acid residues to more than 300. An example of a pro-
tein with multiple domains is shown in Figure 4.21. Note that each domain is a distinct
compact unit consisting of various elements of secondary structure. Domains are usually
connected by loops but they are also bound to each other through weak interactions
formed by the amino acid side chains on the surface of each domain. The top domain of
pyruvate kinase in Figure 4.21 contains residues 1 16 to 219, the central domain contains
residues 1 to 1 15 plus 220 to 388, and the bottom domain contains residues 389 to 530. In
general, domains consist of a contiguous stretch of amino acid residues as in the top and
bottom domains of pyruvate kinase but in some cases a single domain may contain two or
more different regions of the polypeptide chain as in the middle domain.

The evolutionary conservation of protein structure is one of the most important
observations that has emerged from the study of proteins in the past few decades. This
conservation is most easily seen in the case of single-domain homologous proteins from
different species. For example, in Chapter 3 we examined the sequence similarity of cy-
tochrome c and showed that the similarities in primary structure could be used to con-
struct a phylogenetic tree that reveals the evolutionary relationships of the proteins
from different species (Section 3.11). As you might expect, the tertiary structures of cy-
tochrome c proteins are also highly conserved (Figure 4.22). Cytochrome c is an exam-
ple of a protein that contains a heme prosthetic group. The conservation of protein
structure is a reflection of its interaction with heme and its conserved function as an
electron transport protein in diverse species.

Some domain structures occur in many different proteins whereas others are unique. In
general, proteins can be grouped into families according to similarities in domain structures
and amino acid sequence. All of the members of a family have descended from a common
ancestral protein. Some biochemists believe that there may be only a few thousand families

▲ Figure 4.21

Pyruvate kinase from cat ( Felis domesticus).

The main polypeptide chain of this common
enzyme folds into three distinct domains as
indicated by brackets. [PDB 1PKM].

◄ Figure 4.22

Conservation of cytochrome c structure.

(a) Tuna ( Thunnus alalunga ) cytochrome
c bound to heme [PDB 5CYT]. (b) Tuna
cytochrome c polypeptide chain, (c) Rice
( Oryza sativa ) cytochrome c [PDB 1CCR].
(d) Yeast ( Saccharomyces cerevisiae )
cytochrome c [PDB 1YCC]. (e) Bacterial
{Rhodopila globiformis) cytochrome c
[PDB 1HR0].

102 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

(b)

▲ Figure 4.23

Structural similarity of lactate and malate de-
hydrogenase. (a) Bacillus stereothermophilus
lactate dehydrogenase [PDB 1LDN].

(b) Escherichia coli malate dehydrogenase
[PDB 1EMD].

suggesting that all modern proteins are descended from only a few thousand proteins that
were present in the most primitive organisms living 3 billion years ago.

Lactate dehydrogenase and malate dehydrogenase are different enzymes that belong
to the same family of proteins. Their structures are very similar as shown in Figure 4.23.
The sequences of the proteins are only 23% identical. In spite of the obvious similarity
in structure, Nevertheless, this level of sequence similarity is significant enough to con-
clude that the two proteins are homologous. They descend from a common ancestral
gene that duplicated billions of years ago before the last common ancestor of all extant
species of bacteria. Both lactate dehydrogenase and malate dehydrogenase are present in
the same species which is why they are members of a family of related proteins. Protein
families contain related proteins that are present in the same species. The cytochrome c
proteins shown in Figure 4.22 are evolutionarily related but strictly speaking they are
not members of a protein family because there is only one of them in each species. Pro-
tein familes arise from gene duplication events.

Protein domains can be classified by their structures. One commonly used classifi-
cation scheme groups these domains into four categories. The “all- a” category contains
domains that consist almost entirely of a helices and loops. “A11-/3” domains contain only
[3 sheets and nonrepetitive structures that link (3 strands. The other two categories con-
tain domains that have a mixture of a helices and /3 strands. Domains in the u a/f3 ” class
have supersecondary structures such as the (3a(3 motif and others in which regions of
a helix and [3 strand alternate in the polypeptide chain. In the “a + [3 ” category, the do-
mains consist of local clusters of a helices and /3 sheet where each type of secondary
structure arises from separate contiguous regions of the polypeptide chain.

Protein domains can be further classified by the presence of characteristic folds
within each of the four main structural categories. A fold is a combination of secondary
structures that form the core of a domain. Figure 4.24 on pages 103-104 shows selected
examples of proteins from each of the main categories and illustrates a number of com-
mon domain folds. Some domains have easily recognizable folds, such as the / 3 meander
that contains antiparallel [3 strands connected by hairpin loops (Figure 4.20f), or helix
bundles (Figure 4.19c). Other folds are more complex (Figure 4.25).

The important point about Figure 4.24 is not to memorize the structures of com-
mon proteins and folds. The key concept is that proteins can adopt an amazing variety
of different sizes and shapes (tertiary structure) even though they contain only three
basic forms of secondary structure.

The enzymatic activities of lactate
dehydrogenase and malate dehydroge-
nase are compared in Box 7.1.

C. Domain Structure, Function, and Evolution

The relationship between domain structure and function is complex. Often a single do-
main has a particular function such as binding small molecules or catalyzing a single re-
action. In multifunctional enzymes, each catalytic activity can be associated with one of
several domains found in a single polypeptide chain (Figure 4.24j). However, in many
cases the binding of small molecules and the formation of the active site of an enzyme
take place at the interface between two separate domains. These interfaces often form
crevices, grooves, and pockets that are accessible on the surface of the protein. The ex-
tent of contact between domains varies from protein to protein.

The unique shapes of proteins, with their indentations, interdomain interfaces, and
other crevices, allow them to fulfill dynamic functions by selectively and transiently
binding other molecules. This property is best illustrated by the highly specific binding
of reactants (substrates) to substrate -binding sites, or active sites, of enzymes. Because
many binding sites are positioned toward the interior of a protein, they are relatively
free of water. When substrates bind, they fit so well that some of the few remaining
water molecules in the binding site are displaced.

D. Intrinsically Disordered Proteins

This section on tertiary structure wouldn’t be complete without mentioning those pro-
teins and domains that have no stable three-dimensional structure. These intrinsically
disordered proteins (and domains) are quite common and the lack of secondary and
tertiary structure is encoded in the amino acid sequences. There has been selection for

4.8 Quaternary Structure 103

clusters of charged residues (positive or negative) and proline residues that maintain
the polypeptide chain in a disordered state.

Many of these proteins interact with other proteins. They contain short amino acid
sequences that serve as binding sites and these binding sites are within the intrinsically
disordered regions. This allows easy access to the binding site. If a protein contains two
different binding sites for other proteins then the disordered polypeptide chain acts as a
tether to bring the two binding proteins closer together. Several transcription factors
also contain disordered regions when they are not bound to DNA. These regions be-
come ordered when the proteins interact with DNA.

4.8 Quaternary Structure

Many proteins exhibit an additional level of organization called quaternary structure.
Quaternary structure refers to the organization and arrangement of subunits in a pro-
tein with multiple subunits. Each subunit is a separate polypeptide chain. A multisub-
unit protein is referred to as an oligomer (proteins with only one polypeptide chain are
monomers). The subunits of a multisubunit protein may be identical or different.
When the subunits are identical, dimers and tetramers predominate. When the subunits
differ, each type often has a different function. A common shorthand method for de-
scribing oligomeric proteins uses Greek letters to identify types of subunits and sub-
script numerals to indicate numbers of subunits. For example, an cv 2 /3y protein contains
two subunits designated a and one each of subunits designated /3 and y.

The subunits within an oligomeric protein always have a defined stoichiometry and
the arrangement of the subunits gives rise to a stable structure where subunits are usu-
ally held together by weak noncovalent interactions. Hydrophobic interactions are the
principal forces involved although electrostatic forces may contribute to the proper
alignment of the subunits. Because intersubunit forces are usually rather weak, the sub-
units of an oligomeric protein can often be separated in the laboratory. In vivo , however,
the subunits usually remain tightly associated.

Examples of several multisubunit proteins are shown in Figure 4.26. In the case of
triose phosphate isomerase (Figure 4.26a) and HIV protease (Figure 4.26b), the identical
subunits associate through weak interactions between the side chains found mainly in
loop regions. Similar interactions are responsible for the formation of the MS2 capsid
protein that consists of a trimer of identical subunits (Figure 4.26d). In this case, the
trimer units assemble into a more complex structure — the bacteriophage particle. The
enzyme HGPRT (Figure 4.26e) is a tetramer formed from the association of two pairs of
nonidentical subunits. Each of the subunits is a recognizable domain.

The potassium channel protein (Figure 4.26c) is an example of a tetramer of iden-
tical subunits where the subunits interact to form a membrane-spanning region con-
sisting of an eight-helix bundle. The subunits do not form separate domains within the
protein but instead come together to form a single channel. The bacterial photosystem
shown in Figure 4.26f is a complex example of quaternary structure. Three of the sub-
units contribute to a large membrane-bound helix bundle while a fourth subunit (a cy-
tochrome) sits on the exterior surface of the membrane.

Determination of the subunit composition of an oligomeric protein is an essential
step in the physical description of a protein. Typically, the molecular weight of the native
oligomer is estimated by gel- filtration chromatography and then the molecular weight
of each chain is determined by SDS-polyacrylamide gel electrophoresis (Section 3.6).
For a protein having only one type of chain, the ratio of the two values provides the
number of chains per oligomer.

The fact that a large proportion of proteins consist of multiple subunits is probably
related to several factors:

1. Oligomers are usually more stable than their dissociated subunits suggesting that
quaternary structure prolongs the life of a protein in vivo.

2. The active sites of some oligomeric enzymes are formed by residues from adjacent
polypeptide chains.

KEY CONCEPT

There are only three basic types of
secondary structure but thousands of
tertiary folds and domains.

Speculations on the possible relation-
ship between protein domains and
gene organization will be presented
in Chapter 21.

The structures and functions of bacteri-
al and plant photosystems are
described in Chapter 15.

104 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

(c)

E. coli cytochrome b 562

E. coli UDP A/-acetylglucosamine
acyl transferase

Human serum albumin

Human peptidylprolyl
cis/trans isomerase Cow gamma crystallin

Jack bean concanavalin A

Jellyfish green flourescent
protein

▲ Figure 4.24

Examples of tertiary structure in selected proteins, (a) Human {Homo sapiens) serum albumin [PDB 1BJ5] (class: all-a). This protein has several do-
mains consisting of layered a helices and helix bundles, (b) Escherichia coli cytochrome b 5 62 [PDB 1QPU] (class: all-a). This is a heme-binding pro-
tein consisting of a single four-helix bundle domain, (c) Escherichia coli UDP N-acetylglucosamine acyl transferase [PDB 1LXA] (class: a\\-(3). The
structure of this enzyme shows a classic example of a £ helix domain, (d) Jack bean ( Canavalia ensiformis ) concanavalin A [PDB ICON] (class: all -f3).
This carbohydrate-binding protein (lectin) is a single-domain protein made up of a large [3 sandwich fold, (e) Human {Homo sapiens) peptidylprolyl
cis/trans isomerase [PDB 1VBS] (class: a\\-(3). The dominant feature of the structure is a f3 sandwich fold, (f) Cow {Bos taurus) y-crystallin
[PDB 1A45] (class: a 11-/3) This protein contains two (3 barrel domains, (g) Jellyfish {Aequorea victoria) green fluorescent protein [PDB 1GFL] (class:
all -(3). This is a [3 barrel structure with a central a helix. The strands of the sheet are antiparallel, (h) Pig {Sus scrota) retinol-binding protein [PDB
1AQB] (class: a\\-(3). Retinol binds in the interior of a (3 barrel fold. (I) Brewer’s yeast {Saccharomyces carlsburgensis) old yellow enzyme (FMN oxi-
doreductase) [PDB 10YA] (class: alp). The central fold is an al(3 barrel with parallel (3 strands connected by a helices. Two of the connecting a heli-
cal regions are highlighted in yellow, (j) Escherichia colie nzyme required for tryptophan biosynthesis [PDB 1 PI I ] (class: alp). This is a bifunctional
enzyme containing two distinct domains. Each domain is an example of an a/(3 barrel. The left-hand domain contains the indolglycerol phosphate

4.8 Quaternary Structure 105

Yeast FMN oxidoreductase
(old yellow enzyme)

E. coli flavodoxin

Human thioredoxin

Pig adenylyl kinase

E. coli thiol-disulfide
oxidoreductase

E. coli L-arabinose-binding
protein

Neisseria gonorrhea pilin

▲ Figure 4.24 ( continued )

synthetase activity, and the right-hand domain contains the phosphoribosylanthranilate isomerase activity, (k) Pig {Sus scrofa) adenylyl kinase
[PDB 3ADK] (class: alp). This single-domain protein consists of a five-stranded parallel (3 sheet with layers of a helices above and below the sheet.
The substrate binds in the prominent groove between a helices. (I) Escherichia coli flavodoxin [PDB 1AHN] (class: alp). The fold is a five-stranded
parallel twisted sheet surrounded by a helices, (m) Human ( Homo sapiens ) thioredoxin [PDB 1ERU] (class: alp). The structure of this protein is
very similar to that of E. coli flavodoxin except that the five-stranded twisted sheet in the thioredoxin fold contains a single antiparallel strand,
(n) Escherichia coli L-arabinose-binding protein [PDB 1ABE] (class: alp). This is a two-domain protein where each domain is similar to that in E. coli
flavodoxin. The sugar L-arabinose binds in the cavity between the two domains, (o) Escherichia coli DsbA (thiol-disulfide oxidoreductase/disulfide iso-
merase) [PDB 1A23] (class: alp). The predominant feature of this structure is a (mostly) antiparallel (3 sheet sandwiched between a helices. Cysteine
side chains at the end of one of the a helices are shown (sulfur atoms are yellow), (p) Neisseria gonorrhea pilin [PDB 2PIL] (class: a + p). This
polypeptide is one of the subunits of the pili on the surface of the bacteria responsible for gonorrhea. There are two distinct regions of the structure: a
1 3 sheet and a long a helix.

106 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Figure 4.25 ►

Common domain folds.

(a) Parallel twisted sheet

(b) p barrel

3. The three-dimensional structures of many oligomeric proteins change when the pro-
teins bind ligands. Both the tertiary structures of the subunits and the quaternary
structures (i.e., the contacts between subunits) may be altered. Such changes are key
elements in the regulation of the biological activity of certain oligomeric proteins.

4. Different proteins can share the same subunits. Since many subunits have a defined
function (e.g., ligand binding), evolution has favored selection for different combi-
nations of subunits to carry out related functions. This is more efficient than selec-
tion for an entirely new monomeric protein that duplicates part of the function.

5. A multisubunit protein may bring together two sequential enzymatic steps where
the product of the first reaction becomes the substrate of the second reaction. This
gives rise to an effect known as channeling (Section 5.11).

As shown in Figure 4.26, the variety of multisubunit proteins ranges from simple
homodimers such as triose phosphate isomerase to large complexes such as the photo-
systems in bacteria and plants. We would like to know how many proteins are
monomers and how many are oligomers but studies of cell proteomes — the complete
complement of proteins — have only begun.

Table 4.1 on page 108 shows the results of a survey of E. coli proteins in the SWISS-
PROT database. Of those polypeptides that have been analyzed, only about 19% are in
monomers. Dimers are the largest class among the oligomers, and homodimers — where
the two subunits are identical — represent 31% of all proteins. The next largest class is
tetramers of identical subunits. Note that trimers are relatively rare. Most proteins exhibit
dyad symmetry meaning that you can usually draw a line through a protein dividing it
into two halves that are symmetrical about this axis. This dyad symmetry is seen even in

4.8 Quaternary Structure

107

Human hypoxanthine-guanine
phosphoribosyl transferase

Rhodopseudomonas

photosystem

▲ Figure 4.26

Quaternary structure, (a) Chicken {Gallus gal I us) triose phosphate isomerase [PDB 1TIM]. This protein has two identical subunits with a/p barrel folds,
(b) HIV-1 aspartic protease [PDB 1DIF]. This protein has two identical all-/3 subunits that bind symmetrically. HIV protease is the target of many new
drugs designed to treat AIDS patients, (c) Streptomyces lividans potassium channel protein [PDB 1BL8]. This membrane-bound protein has four
identical subunits, each of which contributes to a membrane-spanning eight-helix bundle, (d) Bacteriophage MS2 capsid protein [PDB 2MS2]. The
basic unit of the MS2 capsid is a trimer of identical subunits with a large p sheet, (e) Human ( Homo sapiens) hypoxanthine-guanine phosphoribosyl
transferase (HGPRT) [PDB 1BZY]. HGPRT is a tetrameric protein containing two different types of subunit, (f) Rhodopseudomonas viridis photosys-
tem [PDB 1PRC]. This complex, membrane-bound protein has two identical subunits (orange, blue) and two other subunits (purple, green) bound to
several molecules of photosynthetic pigments.

108 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Table 4.1 Natural occurrence of oligomeric proteins in Escherichia coli

Oligomeric

state

Number of
homooligomers

Number of

heterooligomers Percent

Monomer

19.4

Dimer

115

38.2

Trimer

5.4

Tetramer

21.0

Pentamer

0.1

Hexamer

5.6

Heptamer

0.1

Octamer

2.4

Nonamer

0.0

Decamer

0.0

Undecamer

0.0

Dodecamer

1.6

Higher oligomers

2.2

Polymers

2.7

Figure 4.27 ►

Large protein complexes in the bacterium
Mycoplasma pneumoniae. M. pneumoniae
causes some forms of pneumonia in hu-
mans. This species has one of the smallest
genomes known (689 protein-encoding
genes). Most of those genes are likely to
represent the minimum proteome of a living
cell. The cell contains several large com-
plexes found in all cells: pyruvate dehygro-
genase (purple), ribosome (yellow), GroEL
(red), and RNA polymerase (orange). It also
contains a rod (green) found only in some
bacteria. [Adapted from Kuhner et al. (2009).
Proteome organization in a genome-reduced
bacterium. Science 326:1235-1240]

heterooligomers such as hypoxanthine-guanine phosphoribosyl transferase (HGPRT,
Figure 4.26e) and hemoglobin (Section 4.14). Of course, there are many exceptions, es-
pecially when the oligomers are large complexes.

We will encounter many other examples of multisubunit proteins throughout this
textbook, especially in the chapters on information flow (Chapters 20-22). DNA poly-
merase, RNA polymerase, and the ribosome are excellent examples. Other examples in-
clude GroEL (Section 4.1 ID) and pyruvate dehydrogenase (Section 13.1). Many of
these large proteins are easily seen in electron micrographs, as illustrated in Figure 4.27.

Large complexes are referred to, metaphorically, as protein machines since the vari-
ous polypeptide components work together to carry out a complex reaction. The term

Pyruvate
dehydrogenase
structural core

Ribosome

polymerase

4.9 Protein-Protein Interactions 109

was originally coined to describe complexes such as the replisome (Figure 20.15)
but there are many other examples, including those shown in Figure 4.27.

The bacterial flagellum (Figure 4.28) is a spectacular example of a protein
machine. The complex drives the rotation of a long flagellum using protonmo-
tive force as an energy source (Section 14.3). More than 50 genes are required to
build the flagellum in E. coli but surveys of other bacteria reveal that there are
only about 2 1 core proteins required to build a functional flagellum. The evolu-
tionary history of this protein machine is being actively investigated and it appears
that it was built up by combining simpler components involved in ATP synthesis
and membrane secretion.

4.9 Protein-Protein Interactions

The various subunits in multisubunit proteins bind to each other so strongly that
they rarely dissociate inside the cell. These protein-protein contacts are character-
ized by a number of weak interactions. We have already become familiar with the
type of interactions involved: hydrogen bonds, charge-charge interactions, van der ^

Waals forces, and hydrophobic interactions (Section 2.5). In some cases the contact
areas between two subunits are localized to small patches on the surface of the
polypeptides but while in other cases there can be extensive contact spread over
large portions of the polypeptides. The distinguishing feature of subunit contacts
is the cumulative effect of a large number of individual weak interactions giving a
binding strength that is sufficient to keep the subunits together.

In addition to subunit-subunit contacts, there are many other types of protein-
protein interactions that are less stable. These range from transient contacts between
external proteins and receptors on the cell surface to weak interactions between various
enzymes in metabolic pathways. These weak interactions are much more difficult to detect
but they are essential components of many biochemical reactions.

Consider a simple interaction between two proteins, PI and P2, to give a complex
P1:P2. The equilibrium between the free and bound molecules can be described by either
an association constant (IQ) or a dissociation constant (IQ) (IQ = 1/IQ).

ament cap

F, 9 L ] Hooh-lilament
FWCl iunrtxn

PI + P2 PI :P2

K a =

[PI :P2]

(4.1)

[P1][P2]

PI :P2 PI + P2

Kd =

[P1][P2]

(4.2)

[PI :P2]

FliK 1

FlqO

*[ FlgG [ Dislalrod

-j FlgH | L ring

■j Flgl J P ring

- FliE FlgB

FlgG Proximal rod

F ItF

MS ring

3 FUG

r— FliM

1 C ring

“} ' FUN

FliO

FliP

FliQ

FliR

▲ Figure 4.28

Bacterial flagellum. The bacterial flagellum is
a protein machine composed of 21 core
subunits found in all species (blue boxes).
Two additional subunits are missing in
Firmicutes (white boxes) and five others are
sporadically distributed. The flagellum (hook
+ filament + cap) spins as the motor complex
rotates. The three layers represent the outer
membrane (top), the peptidoglycan layer
(middle), and the cytoplasmic membrane
(bottom). (Courtesy of Howard Ochman.)

Typical association constants for the binding of subunits in a multimeric protein are
greater than 10 8 M -1 (IQ > 10 8 M -1 ) and can range as high as 10 14 M -1 for very tight
interactions. At the other extreme are protein-protein interactions that are so weak they
have no biological significance. These can be fortuitous interactions that arise from time
to time because any two polypeptides will almost always form some kind of weak con-
tact. The lower limit of relevant association constants is about 10 4 M -1 (IQ < 10 4 M -1 ). The
really interesting cases are those with association constants between these two values.

The binding of transcription factors to RNA polymerase is one example of weak
protein-protein interactions that are very important. The association constants range
from about 10 5 M -1 to 10 7 M -1 . The interactions between proteins in signaling pathways
also fall into this range as do the interactions between enzymes in metabolic pathways.

Let’s look at what these association constants mean in terms of protein concen-
trations. As the concentrations of PI and P2 increase it becomes more and more
likely that they will interact and bind to each other. At some concentration, the rate of
binding (a second-order reaction) becomes comparable to the rate of dissociation (a
first-order reaction) and complexes will be present in appreciable amounts. Using the
association constant, we can calculate the ratio of free polypeptide (PI or P2) as a
fraction of the total concentration of either one (PQ or P2 T ). This ratio [free] /[total]
tells us how much of the complex will be present at a given protein concentration.

110 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Figure 4.29 ►

Association constants and protein concentration.

The ratio of free unbound protein to total
protein is shown for a protein-protein inter-
action at three different association constants.
Assuming that the concentration of the other
component is in excess, the concentrations
at which half the molecules are in complex
and half are free corresponds to the recip-
rocal of the association constant. [Adapted
from van Holde, Johnson, and Ho, Principles
of Physical Biochemistry, Prentice Hall.]

[free]

[total]

The curves in Figure 4.29 show these ratios for three different association constants
corresponding to very weak (X a = 10 4 M _1 ), moderate (X a = 10 6 M _1 ), and very strong
(FC a = 10 8 M -1 ) protein-protein interactions. If we assume that one of the components
is present in excess, then the curves represent the concentrations of only the rate-limit-
ing polypeptide. One can demonstrate mathematically that for simple systems the point
at which half of the polypeptide is free and half is in a complex corresponds to the re-
ciprocal of the association constant. For example, if K a = 10 8 M -1 then most of the
polypeptide will be bound at any concentration over 1CT 8 M.

What does this mean in terms of molecules per cell? For an E. coli cell whose
volume is about 2 x 10 -15 1 it means that as long as there are more than a dozen mole-
cules per cell the complex will be stable if K a > 10 8 M _1 . This is why large oligomeric
complexes can exist in E. coli even if there are only a few dozen per cell. Most eukaryotic
cells are 1000 times larger and there must be 12,000 molecules in order to achieve a con-
centration of 10 -8 M. Figure 4.29 also shows why it is impossible for weak interactions
to produce significant numbers of P1:P2 complexes. The protein concentration has to
be greater than 10~ 4 M in order for the complex to be present in significant quantity
and this concentration corresponds to 120,000 molecules in an E. coli cell or 120 million
molecules in a eukaryotic cell. There are no free polypeptides present at such concentra-
tions so weak interactions of this magnitude are biologically meaningless.

There are many techniques for detecting moderate binding. These include direct
techniques such as affinity chromatopraphy, immunoprecipitation, and chemical cross-
linking. Newer techniques rely on more sophisticated manipulations such as phage dis-
play, two-hybrid analysis, and genetic methods. Many workers are attempting to map
the interactions of every protein in the cell using these techniques. An example of such
an “interactome” for many E. coli proteins is shown in Figure 4.30. Note that strong in-
teractions between the subunits of oligomers are easily detected as shown by lines con-
necting the subunits of RNA polymerase, the ribosome, and DNA polymerase. Other
lines connect RNA polymerase to various transcription factors — these represent mod-
erate interactions. Further studies of the “interactome” in various species should give us
a much better picture of the complex protein-protein interactions in living cells.

4.10 Protein Denaturation and Renaturation

Environmental changes or chemical treatments may disrupt the native conformation of
a protein causing loss of biological activity. Such a disruption is called denaturation. The
amount of energy needed to cause denaturation is often small, perhaps equivalent to
that needed for the disruption of three or four hydrogen bonds. Some proteins may unfold
completely when denatured to form a random coil (a fluctuating chain considered to be
totally disordered) but most denatured proteins retain considerable internal structure.
It is sometimes possible to find conditions under which small denatured proteins can
spontaneously renature, or refold, following denaturation.

4.10 Protein Denaturation and Renaturation 111

◄ Figure 4.30

E. coli interactome. Each point on the dia-
gram represents a single E. coli protein. Red
dots are essential proteins and blue dots are
nonessential proteins. Lines joining the
points indicate experimentally determined
protein-protein interactions. Five large com-
plexes are shown: RNA polymerase, DNA
polymerase, ribosome and associated pro-
teins, proteins interacting with cysteine
desulfurase (IscS), and proteins associated
with acyl carrier protein (ACP). (The role of
ACP is described in Section 16.1.)

[Adapted from Butland et al. (2005)]

Proteins are commonly denatured by heating. Under the appropriate conditions,
a modest increase in temperature will result in unfolding and loss of secondary and
tertiary structure. An example of thermal denaturation is shown in Figure 4.31. In this
experiment, a solution containing bovine ribonuclease A is heated slowly and the struc-
ture of the protein is monitored by various techniques that measure changes in confor-
mation. All of these techniques detect a change when denaturation occurs. In the case of
bovine ribonuclease A, thermal denaturation also requires a reducing agent that dis-
rupts internal disulfide bridges allowing the protein to unfold.

Denaturation takes place over a relatively small range of temperature. This indi-
cates that unfolding is a cooperative process where the destabilization of just a few weak
interactions leads to almost complete loss of native conformation. Most proteins have a
characteristic “melting” temperature (T m ) that corresponds to the temperature at the
midpoint of the transition between the native and denatured forms. The T m depends on
pH and the ionic strength of the solution.

Most proteins are stable at temperatures up to 50°C to 60°C under physiological
conditions. Some species of bacteria, such as those that inhabit hot springs and the
vicinity of deep ocean thermal vents, thrive at temperatures well above this range. Pro-
teins in these species denature at much higher temperatures as expected. Biochemists
are actively studying these proteins in order to determine how they resist denaturation.

Proteins can also be denatured by two types of chemicals — chaotropic agents and
detergents (Section 2.4). High concentrations of chaotropic agents, such as urea and
guanidinium salts (Figure 4.32), denature proteins by allowing water molecules to solvate
nonpolar groups in the interior of proteins. The water molecules disrupt the hydrophobic
interactions that normally stabilize the native conformation. The hydrophobic tails of

Figure 4.31 ►

Heat denaturation of ribonuclease A. A solution of ribonuclease A in 0.02 M KOI at pH 2.1 was
heated. Unfolding was monitored by changes in ultraviolet absorbance (blue), viscosity (red), and
optical rotation (green). The y-axis is the fraction of the molecule unfolded at each temperature.
[Adapted from Ginsburg, A., and Carroll, W. R. (1965). Some specific ion effects on the conformation
and thermal stability of ribonuclease. Biochemistry 4:2159-2174.

- i ■ i ■ i ■ I

0 10 20 30 40 50

Temperature (°C)

112 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

h 2 n nh 2

Urea

NH 2 c| ©

h 2 n nh 2

Guanidinium chloride
▲ Figure 4.32

Urea and guanidinium chloride.

▲ Figure 4.33

Disulfide bridges in bovine ribonuclease A. (a) Location of disulfide bridges in the native protein,
(b) View of the disulfide bridge between Cys-26 and Cys-84 [PDB 2AAS].

The numbering convention for amino
acid residues in a polypeptide starts at
the N-terminal end (Section 3.5). Cys-26
is the 26th residue from the N-terminus.

detergents, such as sodium dodecyl sulfate (Figure 2.8), also denature proteins by pene-
trating the protein interior and disrupting hydrophobic interactions.

The native conformation of some proteins (e.g., ribonuclease A) is stabilized by
disulfide bonds. Disulfide bonds are not generally found in intracellular proteins but are
sometimes found in proteins that are secreted from cells. The presence of disulfide bonds
stabilizes proteins by making them less susceptible to unfolding and subsequent degra-
dation when they are exposed to the external environment. Disulfide bond formation
does not drive protein folding; instead, the bonds form where two cysteine residues are
appropriately located once the protein has folded. Formation of a disulfide bond requires
oxidation of the thiol groups of the cysteine residues (Figure 3.4), probably by disulfide-
exchange reactions involving oxidized glutathione, a cysteine -containing tripeptide.

Figure 4.33a shows the locations of the disulfide bridges in ribonuclease A. (Com-
pare this orientation of the protein with that shown in Figure 4.3.) There are four disul-
fide bridges. They can link adjacent (3 strands, (3 strands to a helices, or (3 strands to
loops. Figure 4.33b is a view of the disulfide bridge between a cysteine residue in an
a helix (Cys-26) and a cysteine residue in a (3 strand (Cys-84). Note that the S — S bond
does not align with the cysteine side chains. Disulfide bridges will form whenever the
two cysteine sulfhydryl groups are in close proximity in the native conformation.

Complete denaturation of proteins containing disulfide bonds requires cleavage of
these bonds in addition to disruption of hydrophobic interactions and hydrogen bonds.
2-Mercaptoethanol or other thiol reagents can be added to a denaturing medium in
order to reduce any disulfide bonds to sulfhydryl groups (Figure 4.34). Reduction of the
disulfide bonds of a protein is accompanied by oxidation of the thiol reagent.

In a series of classic experiments, Christian B. Anfinsen and his coworkers studied
the renaturation pathway of ribonuclease A that had been denatured in the presence of
thiol reducing agents. Since ribonuclease A is a relatively small protein (124 amino acid

Figure 4.34 ►

Cleaving disulfide bonds. When a protein
is treated with excess 2-mercaptoethanol
(HSCH 2 CH 2 OH), a disulfide-exchange reac-
tion occurs in which each cystine residue
is reduced to two cysteine residues and
2-mercaptoethanol is oxidized to a disulfide.

H O

W/V N — CH — C WV

H O

w/v N — CH — C WV

2 HSCH 2 CH 2 OH^

s — CH 2 CH 2 OH
s — ch 2 ch 2 oh

WV N — CH — C 'xrx/xr

H O

WV |\| £ |— | £ WV

H O

Cystine residue

Cysteine residues

4.10 Protein Denaturation and Renaturation 113

residues), it refolds (renatures) quickly once it is returned to conditions where the native
form is stable (e.g., cooled below the melting temperature or removed from a solution
containing chaotropic agents). Anfinsen was among the first to show that denatured
proteins can refold spontaneously to their native conformation indicating that the in-
formation required for the native three-dimensional conformation is contained in the
amino acid sequence of the polypeptide chain. In other words, the primary structure
determines the tertiary structure.

Denaturation of ribonuclease A with 8 M urea containing 2-mercaptoethanol re-
sults in complete loss of tertiary structure and enzymatic activity and yields a polypep-
tide chain containing eight sulfhydryl groups (Figure 4.35). When 2-mercaptoethanol is
removed and oxidation is allowed to occur in the presence of urea, the sulfhydryl groups
pair randomly so that only about 1% of the protein population forms the correct four
disulfide bonds recovering original enzymatic activity. (If the eight sulfhydryl groups
pair randomly, 105 disulfide-bonded structures are possible — 7 possible pairings for the
first bond, 5 for the second, 3 for the third, and 1 for the fourth (7x5x3xl = 105) —
but only one of these structures is correct.) However, when urea and 2-mercaptoethanol
are removed simultaneously and dilute solutions of the reduced protein are then exposed
to air, ribonuclease A spontaneously regains its native conformation, its correct set of
disulfide bonds, and its full enzymatic activity. The inactive proteins containing ran-
domly formed disulfide bonds can be renatured if urea is removed, a small amount of 2-
mercaptoethanol is added, and the solution gently warmed. Anfinsens experiments
demonstrate that the correct disulfide bonds can form only after the protein folds into its
native conformation. Anfinsen concluded that the renaturation of ribonuclease A is
spontaneous, driven entirely by the free energy gained in changing to the stable physio-
logical conformation. This conformation is determined by the primary structure.

Proteins occasionally adopt a nonnative conformation and form inappropriate
disulfide bridges when they fold inside a cell. Anfinsen discovered an enzyme, called
protein disulfide isomerase (PDI), that catalyzes reduction of these incorrect bonds. All

▲ Christian B. Anfinsen (1916-1995).

Anfinsen was awarded the Nobel Prize
in Chemistry in 1972 for his work on the
refolding of proteins.

disulfide bonds have been reduced

◄ Figure 4.35

Denaturation and renaturation of ribonuclease A.

Treatment of native ribonuclease A (top) with
urea in the presence of 2-mercaptoethanol
unfolds the protein and disrupts disulfide
bonds to produce reduced, reversibly dena-
tured ribonuclease A (bottom). When the
denatured protein is returned to physiological
conditions in the absence of 2-mercap-
toethanol, it refolds into its native conforma-
tion and the correct disulfide bonds form.
However, when 2-mercaptoethanol alone is
removed, ribonuclease A reoxidizes in the
presence of air, but the disulfide bonds form
randomly, producing inactive protein (such
as the form shown on the right). When urea
is removed, a trace of 2-mercaptoethanol
is added to the randomly reoxidized protein,
and the solution is warmed gently, the disul-
fide bonds break and re-form correctly to
produce native ribonuclease A.

114 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

(a)

Free

energy

Conformation

(b)

▲ Figure 4.36

Energy well of protein folding. The funnels
represent the free-energy potential of folding
proteins, (a) A simplified funnel showing
two possible pathways to the low-energy
native protein. In path B, the polypeptide
enters a local low-energy minimum as it
folds, (b) A more realistic version of the pos-
sible free-energy forms of a folding protein
with many local peaks and dips.

KEY CONCEPT

Most proteins fold spontaneously into a
conformation with the lowest energy.

living cells contain such an activity. The enzyme contains two reduced cysteine residues
positioned in the active site. When the misfolded protein binds, the enzyme catalyzes a
disulfide -exchange reaction whereby the disulfide in the misfolded protein is reduced
and a new disulfide bridge is created between the two cysteine residues in the enzyme.
The misfolded protein is then released and it can refold into the low-energy native
conformation. The structure of the reduced form of E. coli disulfide isomerase (DsbA)
is shown in Figure 4.24o.

4.11 Protein Folding and Stability

New polypeptides are synthesized in the cell by a translation complex that includes
ribosomes, mRNA, and various factors (Chapter 21). As the newly synthesized polypep-
tide emerges from the ribosome, it folds into its characteristic three-dimensional shape.
Folded proteins occupy a low-energy well that makes the native structure much more
stable than alternative conformations (Figure 4.36). The in vitro experiments of Anfmsen
and many other biochemists demonstrate that many proteins can fold spontaneously to
reach this low-energy conformation. In this section we discuss the characteristics of
those proteins that fold into a stable three-dimensional structure.

It is thought that as a protein folds the first few interactions trigger subsequent
interactions. This is an example of cooperative effects in protein folding — the phenom-
enon whereby the formation of one part of a structure leads to the formation of the
remaining parts of the structure. As the protein begins to fold, it adopts lower and lower
energies and begins to fall into the energy well shown in Figure 4.36. The protein may
become temporarily trapped in a local energy well (shown as small dips in the energy
diagram) but eventually it reaches the energy minimum at the bottom of the well. In its
final, stable, conformation, the native protein is much less sensitive to degradation than
an extended, unfolded polypeptide chain. Thus, native proteins can have half-lives of
many cell generations and some molecules may last for decades.

Folding is extremely rapid — in most cases the native conformation is reached in
less than a second. Protein folding and stabilization depend on several noncovalent
forces including the hydrophobic effect, hydrogen bonding, van der Waals interactions,
and charge-charge interactions. Although noncovalent interactions are weak individu-
ally, collectively they account for the stability of the native conformations of proteins.
The weakness of each noncovalent interaction gives proteins the resilience and flexibil-
ity to undergo small conformational changes. (Covalent disulfide bonds also contribute
to the stability of certain proteins.)

In multidomain proteins the different domains fold independently of one another
as much as possible. One of the reasons for limitations on the size of a domain (usually
< 200 residues) is that large domains would fold too slowly if domains were larger than
300 residues. The rate of spontaneous folding would be too slow to be useful.

No actual protein-folding pathway has yet been described in detail but current re-
search is focused on intermediates in the folding pathways of a number of proteins. Sev-
eral hypothetical folding pathways are shown in Figure 4.37. During protein folding, the
polypeptide collapses upon itself due to the hydrophobic effect and elements of second-
ary structure begin to form. This intermediate is called a molten globule. Subsequent
steps involve rearrangement of the backbone chain to form characteristic motifs and, fi-
nally, the stable native conformation.

The mechanism of protein folding is one of the most challenging problems in bio-
chemistry. The process is spontaneous and must be largely determined by the primary
structure (sequence) of the polypeptide. It should be possible, therefore, to predict the
structure of a protein from knowledge of its amino acid sequence. Much progress has
been made in recent years by modeling the folding process using fast computers.

In the remainder of this section, we examine the forces that stabilize protein struc-
ture in more detail. We will also describe the role of chaperones in protein folding.

A. The Hydrophobic Effect

Proteins are more stable in water when their hydrophobic side chains are aggregated in
the protein interior rather than exposed on the surface to the aqueous medium. Because

4.11 Protein Folding and Stability 115

◄ Figure 4.37

Hypothetical protein-folding pathways. The

initially extended polypeptide chains form
partial secondary structures, then approxi-
mate tertiary structures, and finally the
unique native conformations. The arrows
within the structures indicate the direction
from the N- to the C-terminus.

water molecules interact more strongly with each other than with the nonpolar side
chains of a protein, the side chains are forced to associate with one another causing
the polypeptide chain to collapse into a more compact molten globule. The entropy
of the polypeptide decreases as it becomes more ordered. This decrease is more than
offset by the increase in solvent entropy as water molecules that were previously bound
to the protein are released. (Folding also disrupts extended cages of water molecules
surrounding hydrophobic groups.) This overall increase in the entropy of the system
provides the major driving force for protein folding.

Whereas nonpolar side chains are driven into the interior of the protein, most
polar side chains remain in contact with water on the surface of the protein. The sec-
tions of the polar backbone that are forced into the interior of a protein neutralize their
polarity by hydrogen bonding to each other, often generating secondary structures.
Thus, the hydrophobic nature of the interior not only accounts for the association of
hydrophobic residues but also contributes to the stability of helices and sheets. Studies
of folding pathways indicate that hydrophobic collapse and formation of secondary
structures occur simultaneously

Localized examples of this hydrophobic effect are the interactions of the hydropho-
bic side of an amphipathic a helix with the protein core (Section 4.4) and the hy-
drophobic region between (3 sheets in the /3-sandwich structure (Section 4.5). Most of
the examples shown in Figures 4.25 and 4.26 contain juxtaposed regions of secondary
structure that are stabilized by hydrophobic interactions between the side chains of
hydrophobic amino acid residues.

B. Hydrogen Bonding

Hydrogen bonds contribute to the cooperativity of folding and help stabilize the native
conformations of proteins. The hydrogen bonds in a helices, (3 sheets, and turns are the
first to form, giving rise to defined regions of secondary structure. The final native
structure also contains hydrogen bonds between the polypeptide backbone and water,
between the polypeptide backbone and polar side chains, between two polar side
chains, and between polar side chains and water. Table 4.2 shows some of the many
types of hydrogen bonds found in proteins along with their typical bond lengths. Most
hydrogen bonds in proteins are of the N — H — O type. The distance between the donor
and acceptor atoms varies from 0.26 to 0.34 nm and the bonds may deviate from linear-
ity by up to 40°. Recall that hydrogen bonds within the hydrophobic core of a protein
are much more stable than those that form near the surface because the internal hydro-
gen bonds don’t compete with water molecules.

KEY CONCEPT

Entropically driven reactions are reactions
where the most important thermodynamic
change is an increase in entropy of the
system. We can say that the system is
much more disordered at the end of the
reaction than at the beginning. In the case
of hydrophobic interactions, the change
in entropy is mostly due to the release
of ordered water molecules that shield
hydrophobic groups (Section 2.5D).

116

CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Table 4.2 Examples of hydrogen bonds in proteins

Type of

hydrogen bond

Typical distance between
donor and acceptor
atom (nm)

Hydroxyl -hydroxyl

— O— H

— o—
/

0.28

Hydroxyl-carbonyl

— O— H

o = c

0.28

Amide-carbonyl

N— H-

/ \

0.29

Amide-hydroxyl

N— H-

0.30

Amide-imidazole nitrogen

N— H-

r=(

--N^NH

0.31

BOX 4.2 CASP: THE PROTEIN FOLDING GAME

The basic principles of protein folding are reasonably well
understood and it seems certain that if a protein has a sta-
ble three-dimensional structure it will be determined largely
by the primary structure (sequence). This has led to efforts to
predict tertiary structure from knowing the amino acid
sequence. Biochemists have made huge advances in this the-
oretical work in the last 30 years.

The value of such work has to be assessed by making
predictions of the structure of unknown proteins. This led in
1996 to the beginning of CASP-Critical Assessment of
Methods of Protein Structure Prediction. This is a sort of
game with no prizes other than the honor of being success-
ful. Protein folding groups are given the amino acid se-
quences of a number of targets and asked to predict the
three-dimensional structure. The targets are drawn from

those proteins whose structures have just been determined
but the data haven’t yet been published. Contestants have
only a few weeks to send in their predictions before the actual
structures become known.

The results of the 2008 CASP round are shown in the
figure. There were 121 targets and thousands of predictions
were submitted. Success ranged from nearly 100% for easy
proteins to only about 30% for difficult ones. (“Easy” targets
are those where the Protein Data Bank (PDB) already con-
tains the structures of several homologous proteins. “Diffi-
cult” targets are proteins with new folds that have never been
solved.) The success rate for moderately difficult targets has
climbed over the years as the prediction methods improved,
but there’s plenty of opportunity to make winning predic-
tions at the very difficult end of the scale.

Easy

Target difficulty

Difficult

4.11 Protein Folding and Stability

117

C. Van der Waals Interactions and Charge-Charge Interactions

Van der Waals contacts between nonpolar side chains also contribute to the stability of
proteins. The extent of stabilization due to optimized van der Waals interactions is dif-
ficult to determine. The cumulative effect of many van der Waals interactions probably
makes a significant contribution to stability because nonpolar side chains in the interior
of a protein are densely packed.

Charge-charge interactions between oppositely charged side chains may make a small
contribution to the stability of proteins but most ionic side chains are found on the surfaces
where they are solvated and can contribute only minimally to the overall stabilization of
the protein. Nevertheless, two oppositely charged ions occasionally form an ion pair in the
interior of a protein. Such ion pairs are much stronger than those exposed to water.

D. Protein Folding Is Assisted by Molecular Chaperones

Studies of protein folding have led to two general observations regarding the folding of
polypeptide chains into biologically active proteins. First, protein folding does not in-
volve a random search in three-dimensional space for the native conformation. Instead,
protein folding appears to be a cooperative, sequential process in which formation of
the first few structural elements assists in the alignment of subsequent structural fea-
tures. [The need for cooperativity is illustrated by a calculation made by Cyrus
Levinthal. Consider a polypeptide of 100 residues. If each residue had three possible
conformations that could interconvert on a picosecond time scale then a random search
of all possible conformations for the complete polypeptide would take 10 87 seconds —
many times the estimated age of the universe (6 x 10 17 seconds)!]

Second, to a first approximation the folding pattern and the final conformation of a
protein depend on its primary structure. (Many proteins bind metal ions and coenzymes
as described in Chapter 7. These external ligands are also required for proper folding.) As
we saw in the case of ribonuclease A, simple proteins may fold spontaneously into their
native conformations in a test tube without any energy input or assistance. Larger proteins
will also fold spontaneously into their native structures since the final conformation rep-
resents the minimal free energy form. However, larger proteins are more likely to become
temporarily trapped in a local energy well of the type illustrated in Figure 4.36b. The pres-
ence of such metastable incorrect conformations at best slows the rate of protein folding
and at worst causes the folding intermediates to aggregate and fall out of solution. In
order to overcome this problem inside the cell, the rate of correct protein folding is en-
hanced by a group of ubiquitous special proteins called molecular chaperones.

Chaperones increase the rate of correct folding of some proteins by binding newly
synthesized polypeptides before they are completely folded. They prevent the formation
of incorrectly folded intermediates that may trap the polypeptide in an aberrant form.
Chaperones can also bind to unassembled protein subunits to prevent them from ag-
gregating incorrectly and precipitating before they are assembled into a complete multi-
subunit protein.

There are many different chaperones. Most of them are heat shock proteins — pro-
teins that are synthesized in response to temperature increases (heat shock) or other changes
that cause protein denaturation in vivo. The role of heat shock proteins — now recognized
as chaperones — is to repair the damage caused by temperature increases by binding to dena-
tured proteins and helping them to refold rapidly into their native conformation.

The major heat shock protein is Hsp70 (heat shock protein, M r = 70,000). This
protein is present in all species except for some species of archaebacteria. In bacteria, it
is also called DnaK. The normal role of the chaperone Hsp70 is to bind to nascent

► Heat shock proteins. Proteins were synthesized for a short time in the presence of radioactive
amino acids then run on an SDS-polyacrylamide gel. The gel was exposed to film to detect radioactive
proteins. The resulting autoradiograph shows only those proteins that were labeled during the time
of exposure to radioactive amino acids. Lanes “C” are proteins synthesized at normal growth tem-
peratures, and lanes “H” are proteins synthesized during a short heat shock where cells are shifted
to a temperature a few degrees above their normal growth temperature. The induction of heat
shock proteins (chaperones) in four different species is shown. Red dots indicate major heat shock
proteins: top = Hsp90, middle = Hsp70, bottom = Hsp60(GroEL).

i i

C H

mouse

118 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

► Figure 4.38

Escherichia coli chaperonin (GroE). The core
structure consists of two identical rings
composed of seven GroEL subunits. Un-
folded proteins bind to the central cavity.
Bound ATP molecules can be identified by
their red oxygen atoms, (a) Side view, (b)
Top view showing the central cavity. [PDB
1DER]. (c) During folding the size of the
central cavity of one of the rings increases
and the end is capped by a protein contain-
ing seven GroES subunits. [PDB 1AON].

proteins while they are being synthesized in order to prevent aggregation or entrapment
in a local low-energy well. The binding and release of nascent polypeptides is coupled to
the hydrolysis of ATP and usually requires additional accessory proteins. Hsp70/DnaK
is one of the most highly conserved proteins known in all of biology. This indicates that
chaperone-assisted protein folding is an ancient and essential requirement for efficient
synthesis of proteins with the correct three-dimensional structure.

Another important and ubiquitous chaperone is called chaperonin (also called
GroE in bacteria). Chaperonin is also a heat shock protein (Hsp60) that plays an impor-
tant and essential role in assisting normal protein folding inside the cell.

E. coli chaperonin is a complex multisubunit protein. The core structure consists of
two rings containing seven identical GroEL subunits. Each subunit can bind a molecule
of ATP (Figure 4.38a). A simplified version of chaperonin-assisted folding is shown in
Figure 4.39 . Unfolded proteins bind to the hydrophobic central cavity enclosed by the
rings. When folding is complete, the protein is released by hydrolysis of the bound ATP
molecules. The actual pathway is more complicated and requires an additional component
that serves as a cap sealing one end of the central cavity while the folding process takes place.

Figure 4.39 ►

Chaperonin-assisted protein folding. The un-
folded polypeptide enters the central cavity
of chaperonin, where it folds. The hydrolysis
of several ATP molecules is required for
chaperonin function.

Chaperone

4.12 Collagen, a Fibrous Protein

119

The cap contains seven GroES subunits forming an additional ring (Figure 4.38c). The
conformation of the GroEL ring can be altered during folding to increase the size of the
cavity and the role of the cap is to prevent the unfolded protein from being released
prematurely.

As mentioned earlier, some proteins tend to aggregate during folding in the absence
of chaperones. Aggregation is probably due to temporary formation of hydrophobic sur-
faces on folding intermediates. The intermediates bind to each other and the result is that
they are taken out of solution and are no longer able to explore the conformations repre-
sented by the energy funnel shown in Figure 4.36. Chaperonins isolate polypeptide
chains in the folding cavity and thus prevent folding intermediates from aggregating.
The folding cavity serves as an “Anfinsen cage” that allows the chain to reach the correct
low-energy conformation without interference from other folding intermediates.

The central cavity of chaperonin is large enough to accommodate a polypeptide
chain of about 630 amino acid residues (M r = 70,000). Thus, the folding of most small
and medium-sized proteins can be assisted by chaperonin. However, only about 5% to
10% of E. coli proteins (i.e., about 300 different proteins) appear to interact with chap-
eronin during protein synthesis. Medium-sized proteins and those of the a/ (3 structural
class are more likely to require chaperonin-assisted folding. Smaller proteins are able to
fold quickly on their own. Many of the remaining proteins in the cell require other
chaperones, such as HSP70/DnaK.

Chaperones appear to inhibit incorrect folding and assembly pathways by forming
stable complexes with surfaces on polypeptide chains that are exposed only during syn-
thesis, folding, and assembly. Even in the presence of chaperones, protein folding is
spontaneous; for this reason, chaperone-assisted protein folding has been described as
assisted self-assembly.

4.12 Collagen, a Fibrous Protein

To conclude our examination of the three-dimensional structure of proteins, we exam-
ine several proteins to see how their structures are related to their biological functions.
The proteins selected for more detailed study are the structural protein collagen, the
oxygen-binding proteins myoglobin and hemoglobin (Sections 4.12 to 4.13), and anti-
bodies (Section 4.14).

Collagen is the major protein component of the connective tissue of vertebrates. It
makes up about 30% of the total protein in mammals. Collagen molecules have
remarkably diverse forms and functions. For example, collagen in tendons forms stiff,
ropelike fibers of tremendous tensile strength whereas in skin, collagen takes the form
of loosely woven fibers permitting expansion in all directions.

The structure of collagen was worked out by G. N. Ramachandran (famous for his
Ramachandran plots, Section 4.3). The molecule consists of three left-handed heli-
cal chains coiled around each other to form a right-handed supercoil (Figure 4.40).

▲ Figure 4.40

The human type III collagen triple helix. The

extended region of collagen contains three
identical subunits (purple, light blue, and
green). Three left-handed collagen helices
are coiled around one another to form a
right-handed supercoil. [PDB 1BKV]

◄ G.N. Ramachandran (1922-2001). In this
photograph he is illustrating the difference
between an a helix and the left-handed
triple helix of collagen. Note that he has de-
liberately drawn the a helix as a left-handed
helix and not the standard right-handed
form found in most proteins.

120 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

'wv, |\| — Q\-\ — Q 'wv

/ \

h 2 c. .ch 2

X 0H

▲ Figure 4.41

4-Hydroxyproline residue. 4-Hydroxyproline
residues are formed by enzyme-catalyzed
hydroxylation of proline residues.

The requirement for vitamin C is
explained in Section 7.9.

Figure 4.43 ►

5-Hydroxylysine residue. 5-Hydroxylysine
residues are formed by enzyme-catalyzed
hydroxylation of lysine residues.

H?C -

.c.

S N

/h 2 h

'CH 2

◄ Figure 4.42

Interchain hydrogen bonding in collagen. The

amide hydrogen of a glycine residue in one
chain is hydrogen-bonded to the carbonyl
oxygen of a residue, often proline, in an
adjacent chain.

Each left-handed helix in collagen has 3.0 amino acid residues per turn and a pitch of
0.94 nm giving a rise of 0.31 nm per residue. Consequently, a collagen helix is more ex-
tended than an a helix and the coiled-coil structure of collagen is not the same as the
coiled-coil motif discussed in Section 4.7. (Several proteins unrelated to collagen also
form similar three-chain supercoils.)

The collagen triple helix is stabilized by interchain hydrogen bonds. The sequence of
the protein in the helical region consists of multiple repeats of the form -Gly-X-Y-, where
X is often proline and Y is often a modified proline called 4-hydroxyproline (Figure 4.41).
The glycine residues are located along the central axis of the triple helix, where tight pack-
ing of the protein strands can accommodate no other residue. For each -Gly-X-Y- triplet,
one hydrogen bond forms between the amide hydrogen atom of glycine in one chain and
the carbonyl oxygen atom of residue X in an adjacent chain (Figure 4.42). Hydrogen bonds
involving the hydroxyl group of hydroxyproline may also stabilize the collagen triple helix.
Unlike the more common a helix, the collagen helix has no intrachain hydrogen bonds.

In addition to hydroxyproline, collagen contains an additional modified amino
acid residue called 5-hydroxylysine (Figure 4.43). Some hydroxylysine residues are co-
valently bonded to carbohydrate residues, making collagen a glycoprotein. The role of
this glycosylation is not known.

Hydroxyproline and hydroxylysine residues are formed when specific proline and
lysine residues are hydroxylated after incorporation into the polypeptide chains of col-
lagen. The hydroxylation reactions are catalyzed by enzymes and require ascorbic acid
(vitamin C). Hydroxylation is impaired in the absence of vitamin C, and the triple helix
of collagen is not assembled properly.

The limited conformational flexibility of proline and hydroxyproline residues pre-
vents the formation of a helices in collagen chains and also makes collagen somewhat
rigid. (Recall that proline is almost never found in a helices.) The presence of glycine
residues at every third position allows collagen chains to form a tightly wound left-
handed helix that accommodates the proline residues. (Recall that the flexibility of
glycine residues tends to disrupt the right-handed a helix.)

Collagen triple helices aggregate in a staggered fashion to form strong, insoluble
fibers. The strength and rigidity of collagen fibers result in part from covalent

'N — CH — C —

i i

H CH 2

oh 2

CH — OH

T 2

©NH,

4.12 Collagen, a Fibrous Protein 121

( a ) ^

0 = C

\a p 7

CH — CH, — CH,

CH,

<//

+ H,N — CH,

CH,

-ch 2 -

Allysine residue

Lysine residue

C=0

H,0

0 = C C = 0

la /3 7 8 e e 8 7 Pa I

CH — CH 2 — CH 2 — CH 2 — CH = N — CH 2 — CH 2 — CH 2 — CH 2 — CH

(b)

0 = C

Schiff base

H O

\*S

c = o

la P 7 8 s Is 7 Pa I

CH — CH 2 — CH 2 — CH 2 — CH = C — CH 2 — CH 2 — CH

cross-links. The — CH 2 NH 3 + groups of the side chains of some lysine and hydroxyly-
sine residues are converted enzymatically to aldehyde groups ( — CHO), producing ally-
sine and hydroxyallysine residues. Allysine residues (and their hydroxy derivatives) react
with the side chains of lysine and hydroxylysine residues to form Schiff bases, complexes
formed between carbonyl groups and amines (Figure 4.44a). These Schiff bases usually
form between collagen molecules. Allysine residues also react with other allysine
residues by aldol condensation to form cross-links, usually between the individual
strands of the triple helix (Figure 4.44b). Both types of cross-links are converted to
more stable bonds during the maturation of tissues, but the chemistry of these conver-
sions is unknown.

BOX 4.3 STRONGER THAN STEEL

Not all fibrous proteins are composed of a helices. Silk is composed of a number of
proteins that are predominantly / 3 strands. The dragline silk of the spider, Nephila
clavipes , for example, contains two proteins called spidroin 1 and spidroin 2. Both
proteins contain multiple stretches of alanine residues separated by residues that
are mostly glycine. The structure of this silk is not known in spite of major efforts
by many laboratories. However, it is known that the proteins contain extensive
regions of / 3 strands.

There are many different kinds of spider silk and spiders have specialized
glands for each type. The silk fiber produced by the major ampulate gland is
called dragline silk; it is the fiber that spiders use to drop out of danger or anchor
their webs. This silk fiber is quite literally stronger than steel cable. Materials
manufactured from dragline silk would be very useful in a number of applica-
tions, one of which would be personal armor because dragline silk is stronger
than Kevlar. So far it has not been possible to make significant amounts of silk in
the laboratory without relying on spiders.

Nephila clavipes , the golden silk spider. ►

◄ Figure 4.44

Covalent cross-links in collagen, (a) An ally-
sine residue condenses with a lysine residue
to form an intermolecular Schiff-base cross-
link. (b) Two allysine residues condense to
form an intramolecular cross-link.

122 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

▲ Figure 4.45

Chemical structure of the Fe(ll)-protoporphyrin
IX heme group in myoglobin and hemoglobin.

The porphyrin ring provides four of the six
ligands that surround the iron atom.

▲ Figure 4.46

Sperm whale (Physeter catodon) oxymyoglobin.

Myoglobin consists of eight a helices. The
heme prosthetic group binds oxygen (red).
His-64 (green) forms a hydrogen bond with
oxygen, and His-93 (green) is complexed to
the iron atom of the heme. [PDB 1A6M].

▲ John Kendrew’s original model of myoglo-
bin determined from his X-ray diffraction
data in the 1950s. The model is made of
plasticine. It was the first three-dimensional
model of a protein.

4.13 Structures of Myoglobin and Hemoglobin

Like most proteins, myoglobin (Mb) and the related protein hemoglobin (Hb) carry
out their biological functions by selectively and reversibly binding other molecules — in
this case, molecular oxygen (0 2 ). Myoglobin is a relatively small monomeric protein
that facilitates the diffusion of oxygen in vertebrates. It is responsible for supplying oxy-
gen to muscle tissue in reptiles, birds, and mammals. Hemoglobin is a larger tetrameric
protein that carries oxygen in blood.

The red color associated with the oxygenated forms of myoglobin and hemoglobin
(e.g., the red color of oxygenated blood) is due to a heme prosthetic group (Figure 4.45).
(A prosthetic group is a protein-bound organic molecule essential for the activity of the
protein.) Heme consists of a tetrapyrrole ring system (protoporphyrin IX) complexed
with iron. The four pyrrole rings of this system are linked by methene ( — CH=)
bridges so that the unsaturated porphyrin is highly conjugated and planar. The bound
iron is in the ferrous, or Fe®, oxidation state; it forms a complex with six ligands, four
of which are the nitrogen atoms of protoporphyrin IX. (Other proteins, such as cy-
tochrome a and cytochrome c, contain different porphyrin/heme groups.)

Myoglobin is a member of a family of proteins called globins. The tertiary structure
of sperm whale myoglobin shows that the protein consists of a bundle of eight a helices
(Figure 4.46). It is a member of the all -a structural category. The globin fold has several
groups of a helices that form a layered structure. Adjacent helices in each layer are tilted
at an angle that allows the side chains of the amino acid residues to interdigitate.

The interior of myoglobin is made up almost exclusively of hydrophobic amino
acid residues, particularly those that are highly hydrophobic — valine, leucine, isoleucine,
phenylalanine, and methionine. The surface of the protein contains both hydrophilic
and hydrophobic residues. As is the case with most proteins, the tertiary structure of
myoglobin is stabilized by hydrophobic interactions within the core. Folding of the
polypeptide chain is driven by the energy minimization that results from formation of
this hydrophobic core.

The heme prosthetic group of myoglobin occupies a hydrophobic cleft formed by
three a helices and two loops. The binding of the porphyrin moiety to the polypeptide is
due to a number of weak interactions including hydrophobic interactions, van der Waals
contacts, and hydrogen bonds. There are no covalent bonds between the porphyrin and
the amino acid side chains of myoglobin. The iron atom of heme is the site of oxygen
binding as shown in Figure 4.46. Two histidine residues interact with the iron atom and
the bound oxygen. Accessibility of the heme group to molecular oxygen depends on
slight movement of nearby amino acid side chains. We will see later that the hydrophobic
crevices of myoglobin and hemoglobin are essential for the reversible binding of oxygen.

In vertebrates, 0 2 is bound to molecules of hemoglobin for transport in red blood
cells, or erythrocytes. Viewed under a microscope, a mature mammalian erythrocyte is a
biconcave disk that lacks a nucleus or other internal membrane-enclosed compart-
ments (Figure 4.47). A typical human erythrocyte is filled with approximately 3 X 10 8
hemoglobin molecules.

Hemoglobin is more complex than myoglobin because it is a multisubunit protein.
In adult mammals, hemoglobin contains two different globin subunits called a-globin
and (3-globin . Hemoglobin is an a 2 /3 2 tetramer — it contains two a chains and two
/ 3 chains. Each of these globin subunits is similar in structure and sequence to myoglobin
reflecting their evolution from a common ancestral globin gene in primitive chordates.

Each of the four globin subunits contains a heme prosthetic group identical to that
found in myoglobin. The a and [3 subunits face each other across a central cavity
(Figure 4.48). The tertiary structure of each of the four chains is almost identical to that
of myoglobin (Figure 4.49). The a chain has seven a helices, and the [3 chain has eight.
(Two short a helices found in (3 - globin and myoglobin are fused into one larger one in
a-globin) Hemoglobin, however, is not simply a tetramer of myoglobin molecules. Each
a chain interacts extensively with a / 3 chain so hemoglobin is actually a dimer of a(3 sub-
units. We will see in the following section that the presence of multiple subunits is respon-
sible for oxygen-binding properties that are not possible with single- chain myoglobin.

4.14 Oxygen Binding to Myoglobin and Hemoglobin 123

▲ Figure 4.48

Human {Homo sapiens) oxyhemoglobin, (a) Structure of human oxyhemoglobin showing two a and two
/3 subunits. Heme groups are shown as stick models. [PDB 1HND]. (b) Schematic diagram of the
hemoglobin tetramer. The heme groups are red.

4.14 Oxygen Binding to Myoglobin and Hemoglobin

The oxygen-binding activities of myoglobin and hemoglobin provide an excellent ex-
ample of how protein structure relates to physiological function. These proteins are
among the most intensely studied proteins in biochemistry. They were the first complex
proteins whose structure was determined by X-ray crystallography (Section 4.2). A
number of the principles described here for oxygen-binding proteins also hold true for
the enzymes that we will study in Chapters 5 and 6. In this section we examine the
chemistry of oxygen binding to heme, the physiology of oxygen binding to myoglobin
and hemoglobin, and the regulatory properties of hemoglobin.

A. Oxygen Binds Reversibly to Heme

We will use myoglobin as an example of oxygen binding to the heme prosthetic group
but the same principles apply to hemoglobin. The reversible binding of oxygen is called
oxygenation. Oxygen- free myoglobin is called deoxy myoglobin and the oxygen-bearing
molecule is called oxymyoglobin. (The two forms of hemoglobin are called deoxyhemoglobin
and oxyhemoglobin.)

Some substituents of the heme prosthetic group are hydrophobic — this feature
allows the prosthetic group to be partially buried in the hydrophobic interior of the
myoglobin molecule. Recall from Figure 4.46 that there are two polar residues, His-64
and His -93, situated near the heme group. In oxymyoglobin, six ligands are coordinated
to the ferrous iron, with the ligands in octahedral geometry around the metal cation
(Figures 4.50 and 4.51). Four of the ligands are the nitrogen atoms of the tetrapyrrole ring
system; the fifth ligand is an imidazole nitrogen from His- 93 (called the proximal histidine);
and the sixth ligand is molecular oxygen bound between the iron and the imidazole side
chain of His-64 (called the distal histidine). In deoxymyoglobin, the iron is coordinated to
only five ligands because oxygen is not present. The nonpolar side chains of Val-68 and
Phe-43, shown in Figure 4.51, contribute to the hydrophobicity of the oxygen-binding
pocket and help hold the heme group in place. Several side chains block the entrance to the
heme-containing pocket in both oxymyoglobin and deoxymyoglobin. The protein struc-
ture in this region must vibrate, or breathe, rapidly to allow oxygen to bind and dissociate.

The hydrophobic crevice of the globin polypeptide holds the key to the ability of myo-
globin and hemoglobin to suitably bind and release oxygen. Free heme does not reversibly
bind oxygen in aqueous solution; instead, the Fe© of the heme is almost instantly ox-
idized to Fe©. (Oxidation is equivalent to the loss

of an electron, as described in Section 6. 1C. Reduction is the gain of an electron. Oxida-
tion and reduction refer to the transfer of electrons and not to the presence or absence of
oxygen molecules.)

v Figure 4.47

Scanning electron micrograph of mammalian
erythrocytes. Each cell contains approxi-
mately 300 million hemoglobin molecules.
The cells have been artificially colored.

▲ Figure 4.49

Tertiary structure of myoglobin, a-globin, and
/Fglobin. The orientations of the individual
a-globin and /3-globin subunits of hemoglo-
bin have been shifted in order to reveal the
similarities in tertiary structure. The three
structures have been superimposed. All
of the structures are from the oxygenated
forms shown in Figures 4.46 and 4.48.
Color code: a-globin (blue), /3-globin
(purple), myoglobin (green).

124 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

His-64

Heme

▲ Figure 4.50

Oxygen-binding site of sperm whale oxymyo-
globin. The heme prosthetic group is repre-
sented by a parallelogram with a nitrogen
atom at each corner. The blue dashed lines
illustrate the octahedral geometry of the
coordination complex.

▲ Figure 4.51

The oxygen-binding site in sperm whale myo-
globin. Fed I) (orange) lies in the plane of
the heme group. Oxygen (green) is bound to
the iron atom and the amino acid side chain
of His-64. Val-68 and Phe-43 contribute to
the hydrophobic environment of the oxygen-
binding site. [PDB 1AGM].

The structure of myoglobin and hemoglobin prevents the permanent transfer of an
electron or irreversible oxidation thereby ensuring the reversible binding of molecular
oxygen for transport. The ferrous iron atom of heme in hemoglobin is partially oxi-
dized when 0 2 is bound. An electron is temporarily transferred toward the oxygen atom
that is attached to the iron so that the molecule of dioxygen is partially reduced. If the
electron were transferred completely to the oxygen, the complex would be Fe 3+ — 0 2 ®
(a superoxide anion attached to ferric iron). The globin crevice prevents complete elec-
tron transfer and enforces return of the electron to the iron atom when 0 2 dissociates.

B. Oxygen-Binding Curves of Myoglobin and Hemoglobin

Oxygen binds reversibly to myoglobin and hemoglobin. The extent of binding at equi-
librium depends on the concentration of the protein and the concentration of oxygen.
This relationship is depicted in oxygen-binding curves (Figure 4.52). In these figures,
the fractional saturation ( Y ) of a fixed amount of protein is plotted against the concen-
tration of oxygen (measured as the partial pressure of gaseous oxygen, p0 2 ). The frac-
tional saturation of myoglobin or hemoglobin is the fraction of the total number of
molecules that are oxygenated.

[Mb0 2 ]

[Mb0 2 ] + [Mb]

(4.3)

The oxygen-binding curve of myoglobin is hyperbolic (Figure 4.52), indicating that there
is a single equilibrium constant for the binding of 0 2 to the macromolecule. In con-
trast, the curve depicting the relationship between oxygen concentrations and binding
to hemoglobin is sigmoidal. Sigmoidal (S-shaped) binding curves indicate that more
than one molecule of ligand is binding to each protein. In this case, up to four mole-
cules of 0 2 bind to hemoglobin, one per heme group of the tetrameric protein. The
shape of the curve indicates that the oxygen-binding sites of hemoglobin interact such
that the binding of one molecule of oxygen to one heme group facilitates binding of
oxygen molecules to the other hemes. The oxygen affinity of hemoglobin increases as
each oxygen molecule is bound. This interactive binding phenomenon is termed
positive cooperativity of binding.

The partial pressure at half- saturation (P 50 ) is a measure of the affinity of the pro-
tein for 0 2 . A low P 50 indicates a high affinity for oxygen since the protein is half-satu-
rated with oxygen at a low oxygen concentration; similarly, a high P 50 signifies a low
affinity. Myoglobin molecules are half- saturated at a p0 2 of 2.8 torr (1 atmosphere =
760 torr). The P 50 for hemoglobin is much higher (26 torr) reflecting its lower affinity
for oxygen. The heme prosthetic groups of myoglobin and hemoglobin are identical but
the affinities of these groups for oxygen differ because the microenvironments provided
by the proteins are slightly different. Oxygen affinity is an intrinsic property of the pro-
tein. It is similar to the equilibrium binding/dissociation constants that are commonly
used to describe the binding of ligands to other proteins and enzymes (Section 4.9).

As Figure 4.52 shows, at the highp0 2 found in the lungs (about 100 torr) both myo-
globin and hemoglobin are nearly saturated. However, at p0 2 values below about 50 torr,
myoglobin is still almost fully saturated whereas hemoglobin is only partially saturated.
Much of the oxygen carried by hemoglobin in erythrocytes is released within the capillar-
ies of tissues where p0 2 is low (20 to 40 torr). Myoglobin in muscle tissue then binds oxy-
gen released from hemoglobin. The differential affinities of myoglobin and hemoglobin
for oxygen thus lead to an efficient system for oxygen delivery from the lungs to muscle.

The cooperative binding of oxygen by hemoglobin can be related to changes in the
protein conformation that occur on oxygenation. Deoxyhemoglobin is stabilized by
several intra- and intersubunit ion pairs. When oxygen binds to one of the subunits,
it causes a movement that disrupts these ion pairs and favors a slightly different conforma-
tion. The movement is triggered by the reactivity of the heme iron atom (Figure 4.53).
In deoxyhemoglobin, the iron atom is bound to only five ligands (as in myoglobin). It is
slightly larger than the cavity within the porphyrin ring and lies below the plane of the ring.
When 0 2 — the sixth ligand — binds to the iron atom, the electronic structure of the iron

4.14 Oxygen Binding to Myoglobin and Hemoglobin 125

(a) (b)

p0 2 (torr) p0 2 (torr)

▲ Figure 4.52

Oxygen-binding curves of myoglobin and hemoglobin, (a) Comparison of myoglobin and hemoglobin. The fractional saturation (VO of each protein is
plotted against the partial pressure of oxygen (p02). The oxygen-binding curve of myoglobin is hyperbolic, with half-saturation {Y = 0.5) at an oxygen
pressure of 2.8 torr. The oxygen-binding curve of hemoglobin in whole blood is sigmoidal, with half-saturation at an oxygen pressure of 26 torr.
Myoglobin has a greater affinity than hemoglobin for oxygen at all oxygen pressures. In the lungs, where the partial pressure of oxygen is high, hemo-
globin is nearly saturated with oxygen. In tissues, where the partial pressure of oxygen is low, oxygen is released from oxygenated hemoglobin and
transferred to myoglobin, (b) O 2 binding by the different states of hemoglobin. The oxy (R, or high-affinity) state of hemoglobin has a hyperbolic
binding curve. The deoxy (T, or low-affinity) state of hemoglobin would also have a hyperbolic binding curve but with a much higher concentration for
half-saturation. Solutions of hemoglobin containing mixtures of low- and high-affinity forms show sigmoidal binding curves with intermediate oxygen
affinities.

Porphyrin plane Fe

▲ Figure 4.53

Conformational changes in a hemoglobin chain induced by oxygenation. When the heme iron of a he-
moglobin subunit is oxygenated (red), the proximal histidine residue is pulled toward the porphyrin
ring. The helix containing the histidine also shifts position, disrupting ion pairs that cross-link the
subunits of deoxyhemoglobin (blue).

126

CHAPTER 4 Proteins: Three-Dimensional Structure and Function

changes, its diameter decreases, and it moves into the plane of the porphyrin ring
pulling the helix that contains the proximal histidine. The change in tertiary structure
results in a slight change in quaternary structure and this allows the remaining subunits
to bind oxygen more readily. The entire tetramer appears to shift from the deoxy to the
oxy conformation only after at least one oxygen molecule binds to each a(3 dimer. (For
further discussion, see Section 5.9C.)

The conformational change of hemoglobin is responsible for the positive cooperativ-
ity of binding seen in the binding curve (Figure 4.52a). The shape of the curve is due to
the combined effect of the two conformations (Figure 4.52b). The completely deoxy-
genated form of hemoglobin has a low affinity for oxygen and thus exhibits a hyperbolic
binding curve with a very high concentration of half- saturation. Only a small amount of
hemoglobin is saturated at low oxygen concentrations. As the concentration of oxygen in-
creases, some of the hemoglobin molecules bind a molecule of oxygen and this increases
their affinity for oxygen so that they are more likely to bind additional oxygen. This causes
the sigmoidal curve and also a sharp rise in binding. More molecules of hemoglobin are in
the oxy conformation. If all of the hemoglobin molecules were in the oxy conformation, a
solution would exhibit a hyperbolic binding curve. Release of the oxygen molecules allows
the hemoglobin molecule to re-form the ion pairs and resume the deoxy conformation.

The two conformations of hemoglobin are called the T (tense) and R (relaxed)
states, using the standard terminology for such conformational changes. In hemoglo-
bin, the deoxy conformation, which resists oxygen binding, is considered the inactive
(T) state, and the oxy conformation, which facilitates oxygen binding, is considered the
active (R) state. The R and T states are in dynamic equilibrium.

BOX 4.4 EMBRYONIC AND FETAL HEMOGLOBINS

The human a globin genes are located on chromosome 16 in
a cluster of related members of the globin gene family. There
are two different genes encoding a globin: aq and a 2 Up-
stream of these genes there is another functional gene called
£ (zeta). The locus includes two nonfunctional pseudogenes,
one related to £ and the other derived from a duplicated
a globin gene

The f3 globin gene is on chromosome 1 1 and it is also lo-
cated at a locus where there are other members of the globin
gene family. The functional genes are d, two related y globin
genes (y A and y G ), and an s (epsilon) gene. This locus also
contains a pseudogene related to [3 (if/p).

The other globin genes encode hemoglobin subunits that
are expressed in the early embryo and in the fetus. The embry-
onic hemoglobins are called Gower 1 (£ 2 s 2 ), Gower 2 (a 2 s 2 ),
and Portland (£ 272 )- The fetal hemoglobin has the subunit
composition a 2 y 2 . The adult hemoglobins are a 2 f 3 2 and a 2 8 2 .

During early embryogenesis, the growing embryo gets
oxygen from the mother’s blood through the placenta.
The concentration of oxygen in the embryo is much lower
than the concentration of oxygen in adult blood. The embry-

onic hemoglobins compensate by binding oxygen much more
tightly, their P 50 values range from 4 to 12 torr — much lower
than the value of adult hemoglobin (26 torr). The fetal hemo-
globins bind oxygen less tightly than the embryonic hemoglo-
bin but tighter than the adult hemoglobins (P 50 = 20 torr).

Expression of the various globin genes is carefully regu-
lated so that the right genes are transcribed at the right time.
Sometimes mutations arise where the fetal y globin genes are
inappropriately expressed in adults. The result is a phenotype
known as Hereditary Persistence of Fetal Hemoglobin
(HPFH). This is just one of hundreds of hemoglobin variants
that have been detected in humans. You can read about them
on a database called Online Mendelian Inheritance in Man
(OMIM), the most complete and accurate database of
human genetic diseases (ncbi.nlm.nih.gov/omim).

► Human fetus.

Chromosome 16

◄ Globin genes,
cq a 2

y G ? A

s P

Chromosome 1 1

4.14 Oxygen Binding to Myoglobin and Hemoglobin 127

▲ Julian Voss-Andreae created a sculpture called “Heart of Steel (Hemoglobin)” in 2005 in the City of Lake Oswego, Oregon. The sculpture is a
depiction of a hemoglobin molecule with a bound oxygen atom. The original sculpture was shiny steel (left). After 10 days (middle) it had started
to rust as the iron in the steel reacted with oxygen in the atmosphere. After several months (right) the sculpture was completely rust colored.

C. Hemoglobin Is an Allosteric Protein

The binding and release of oxygen by hemoglobin are regulated by allosteric interactions
(from the Greek alios , “other”). In this respect, hemoglobin — a carrier protein, not an
enzyme — resembles certain regulatory enzymes (Section 5.9). Allosteric interactions
occur when a specific small molecule, called an allosteric modulator, or allosteric effector,
binds to a protein (usually an enzyme) and modulates its activity. The allosteric modu-
lator binds reversibly at a site separate from the functional binding site of the protein.
An effector molecule may be an activator or an inhibitor. A protein whose activity is
modulated by allosteric effectors is called an allosteric protein.

Allosteric modulation is accomplished by small but significant changes in the con-
formations of allosteric proteins. It involves cooperativity of binding that is regulated by
binding of the allosteric effector to a distinct site that doesn’t overlap the normal bind-
ing site of a substrate, product, or transported molecule such as oxygen. An allosteric
protein is in an equilibrium in which its active shape (R state) and its inactive shape
(T state) are rapidly interconverting. A substrate, which obviously binds at the active
site (to heme in hemoglobin), binds most avidly when the protein is in the R state. An
allosteric inhibitor, which binds at an allosteric or regulatory site, binds most avidly to
the T state. The binding of an allosteric inhibitor to its own site causes the allosteric
protein to change rapidly from the R state to the T state. The binding of a substrate to
the active site (or an allosteric activator to the allosteric site) causes the reverse change.
The change in conformation of an allosteric protein caused by binding or release of an
effector extends from the allosteric site to the functional binding site (the active site).
The activity level of an allosteric protein depends on the relative proportions of mole-
cules in the R and T forms and these, in turn, depend on the relative concentrations of
the substrates and modulators that bind to each form.

The molecule 2,3-frisphospho-D-glycerate (2,3BPG) is an allosteric effector of
mammalian hemoglobin. The presence of 2,3BPG in erythrocytes raises the P 50 for
binding of oxygen to adult hemoglobin to about 26 torr — much higher than the P 50 for
oxygen binding to purified hemoglobin in aqueous solution (about 12 torr). In other
words, 2,3BPG in erythrocytes substantially lowers the affinity of deoxyhemoglobin for
oxygen. The concentrations of 2,3BPG and hemoglobin within erythrocytes are nearly
equal (about 4.7 mM).

H — C — OPO

1 ©

ch 2 opo 3 ^

▲ 2,3-Bisphospho-D-glycerate (2,3BPG).

The synthesis of 2,3BPG in red blood
cells is described in Box 11.2
(Chapter 11).

128 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

Figure 4.54 ►

Binding of 2,3BPG to deoxyhemoglobin. The

central cavity of deoxyhemoglobin is lined
with positively charged groups that are com-
plementary to the carboxylate and phos-
phate groups of 2,3BPG. Both 2,3BPG and
the ion pairs shown help stabilize the deoxy
conformation. The a subunits are shown in
pink, the (3 subunits in blue, and the heme
prosthetic groups in red.

R and T conformations are explained
more thoroughly in Section 5.10,
“Theory of Allostery.”

▲ Figure 4.55

Bohr effect. Lowering the pH decreases the
affinity of hemoglobin for oxygen.

The effector 2,3BPG binds in the central cavity of hemoglobin between the two
[3 subunits. In this binding pocket there are six positively charged side chains and the
N-terminal a-amino group of each / 3 chain forming a cationic binding site (Figure 4.54).
In deoxyhemoglobin, these positively charged groups can interact electrostatically with
the five negative charges of 2,3BPG. When 2,3BPG is bound, the deoxy conformation
(the T state, which has a low affinity for 0 2 ) is stabilized and conversion to the oxy con-
formation (the R or high-affinity state) is inhibited. In oxyhemoglobin, the [3 chains are
closer together and the allosteric binding site is too small to accommodate 2,3BPG. The
reversibly bound ligands 0 2 and 2,3BPG have opposite effects on the R T equilib-
rium. Oxygen binding increases the proportion of hemoglobin molecules in the oxy (R)
conformation and 2,3BPG binding increases the proportion of hemoglobin molecules
in the deoxy (T) conformation. Because oxygen and 2,3BPG have different binding
sites, 2,3BPG is a true allosteric effector.

In the absence of 2,3BPG, hemoglobin is nearly saturated at an oxygen pressure of
about 20 torn Thus, at the low partial pressure of oxygen that prevails in the tissues (20 to
40 torr), hemoglobin without 2,3BPG would not unload its oxygen. In the presence of
equimolar 2,3BPG, however, hemoglobin is only about one-third saturated at 20 torr. The
allosteric effect of 2,3BPG causes hemoglobin to release oxygen at the low partial pressures
of oxygen in the tissues. In muscle, myoglobin can bind some of the oxygen that is released.

Additional regulation of the binding of oxygen to hemoglobin involves carbon
dioxide and protons, both of which are products of aerobic metabolism. C0 2 decreases
the affinity of hemoglobin for 0 2 by lowering the pH inside red blood cells. Enzyme-
catalyzed hydration of C0 2 in erythrocytes produces carbonic acid, H 2 C0 3 , which dis-
sociates to form bicarbonate and a proton thereby lowering the pH.

C0 2 + H 2 0 H 2 C0 3 H© + HC0 3 © (4.4)

The lower pH leads to protonation of several groups in hemoglobin. These groups then
form ion pairs that help stabilize the deoxy conformation. The increase in the concentration
of C0 2 and the concomitant decrease in pH raise the P 50 of hemoglobin (Figure 4.55).
This phenomenon, called the Bohr effect, increases the efficiency of the oxygen delivery
system. In inhaling lungs, where the C0 2 level is low, 0 2 is readily picked up by
hemoglobin; in metabolizing tissues, where the C0 2 level is relatively high and the pH is
relatively low, 0 2 is readily unloaded from oxyhemoglobin.

4.15 Antibodies Bind Specific Antigens 129

Carbon dioxide is transported from the tissues to the lungs in two ways. Most C0 2
produced by metabolism is transported as dissolved bicarbonate ions but some carbon
dioxide is carried by hemoglobin itself the form of carbamate adducts (Figure 4.56). At
the pH of red blood cells (7.2) and at high concentrations of C0 2 , the unprotonated
amino groups of the four N- terminal residues of deoxyhemoglobin (pFC a values between 7
and 8) can react reversibly with C0 2 to form carbamate adducts. The carbamates of oxy-
hemoglobin are less stable than those of deoxyhemoglobin. When hemoglobin reaches
the lungs, where the partial pressure of C0 2 is low and the partial pressure of 0 2 is high,
hemoglobin is converted to its oxygenated state and the C0 2 that was bound is released.

4.15 Antibodies Bind Specific Antigens

Vertebrates possess a complex immune system that eliminates foreign substances includ-
ing infectious bacteria and viruses. As part of this defense system, vertebrates synthesize
proteins called antibodies (also known as immunoglobulins) that specifically recognize
and bind antigens. Many different types of foreign compounds can serve as antigens that
produce an immune response. Antibodies are synthesized by white blood cells called
lymphocytes — each lymphocyte and its descendants synthesize the same antibody. Be-
cause animals are exposed to many foreign substances over their lifetimes, they develop a
huge array of antibody-producing lymphocytes that persist at low levels for many years
and can later respond to the antigen during reinfection. The memory of the immune sys-
tem is the reason certain infections do not recur in an individual despite repeated expo-
sure. Vaccines (inactivated pathogens or analogs of toxins) administered to children are
effective because immunity established in childhood lasts through adulthood.

When an antigen — either novel or previously encountered — binds to the surface of
lymphocytes, these cells are stimulated to proliferate and produce soluble antibodies for
secretion into the bloodstream. The soluble antibodies bind to the foreign organism or
substance forming antibody-antigen complexes that precipitate and mark the antigen
for destruction by a series of interacting proteases or by lymphocytes that engulf the
antigen and digest it intracellularly.

The most abundant antibodies in the bloodstream are of the immunoglobulin
G class (IgG). These are Y-shaped oligomers composed of two identical light chains and
two identical heavy chains connected by disulfide bonds (Figure 4.57). Immunoglobulins
are glycoproteins containing covalently bound carbohydrates attached to the heavy
chains. The N-termini of pairs of light and heavy chains are close together. Light chains contain
two domains and heavy chains contain four domains. Each of the domains consists of

o —

c — N — R

▲ Figure 4.56

Carbamate adduct. Carbon dioxide produced
by metabolizing tissues can react reversibly
with the N-terminal residues of the globin
chains of hemoglobin, converting them to
carbamate adducts.

(a)

(b)

Antigen-binding

site

Antigen-binding

site

◄ Figure 4.57

Human antibody structure, (a) Structure.

I I (b) Diagram. Two heavy chains (blue) and

two light chains (red) of antibodies of the
immunoglobulin G class are joined by disul-
fide bonds (yellow). The variable domains of
I | both the light and heavy chains (where

®OOC COO® antigen binds) are colored more darkly.

130 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

▲ Figure 4.58

The immunoglobulin fold. The domain con-
sists of a sandwich of two antiparallel
/ 3 sheets. [PDB 1REI].

about 110 residues assembled into a common motif called the immunoglobulin fold whose
characteristic feature is a sandwich composed of two antiparallel /3 sheets (Figure 4.58).
This domain structure is found in many other proteins of the immune system.

The N-terminal domains of antibodies are called the variable domains because
of their sequence diversity. They determine the specificity of antigen binding. X-ray
crystallographic studies have shown that the antigen-binding site of a variable do-
main consists of three loops, called hypervariable regions, that differ widely in size
and sequence. The loops from a light chain and a heavy chain combine to form a
barrel, the upper surface of which is complementary to the shape and polarity of a
specific antigen. The match between the antigen and antibody is so close that there
is no space for water molecules between the two. The forces that stabilize the inter-
action of antigen with antibody are primarily hydrogen bonds and electrostatic in-
teractions. An example of the interaction of antibodies with a protein antigen is
shown in Figure 4.59.

Antibodies are used in the laboratory for the detection of small quantities of vari-
ous substances because of their remarkable antigen-binding specificity. In a common
type of immunoassay, fluid containing an unknown amount of antigen is mixed with a
solution of labeled antibody and the amount of antibody-antigen complex formed is
measured. The sensitivity of these assays can be enhanced in a variety of ways to make
them suitable for diagnostic tests.

▲ Figure 4.59

Binding of three different antibodies to an antigen (the protein lysozyme). The structures of the three
antigen-antibody complexes have been determined by X-ray crystallography. This composite view,
in which the antigen and antibodies have been separated, shows the surfaces of the antigen and
antibodies that interact. Only parts of the three antibodies are shown.

Summary

1. Proteins fold into many different shapes, or conformations. Many
proteins are water-soluble, roughly spherical, and tightly folded.
Others form long filaments that provide mechanical support to
cells and tissues. Membrane proteins are integral components of
membranes or are associated with membranes.

2. There are four levels of protein structure: primary (sequence of
amino acid residues), secondary (regular local conformation,
stabilized by hydrogen bonds), tertiary (compacted shape of the
entire polypeptide chain), and quaternary (assembly of two or
more polypeptide chains into a multisubunit protein).

3. The three-dimensional structures of biopolymers, such as
proteins can be determined by X-ray crystallography and NMR
spectroscopy.

4. The peptide group is polar and planar. Rotation around the
N — C a and C a — C bonds is described by <p and if/.

5. The a helix, a common secondary structure, is a coil containing
approximately 3.6 amino acid residues per turn. Hydrogen bonds
between amide hydrogens and carbonyl oxygens are roughly par-
allel to the helix axis.

Problems 131

6. The other common type of secondary structure, /3 structure,
often consists of either parallel or antiparallel /3 strands that are
hydrogen-bonded to each other to form /3 sheets.

7. Most proteins include stretches of nonrepeating conformation,
including turns and loops that connect a helices and (3 strands.

8. Recognizable combinations of secondary structural elements are
called motifs.

9. The tertiary structure of proteins consists of one or more do-
mains, which may have recognizable structures and may be asso-
ciated with particular functions.

10 . In proteins that possess quaternary structure, subunits are usually
held together by noncovalent interactions.

11. The native conformation of a protein can be disrupted by the ad-
dition of denaturing agents. Renaturation may be possible under
certain conditions.

12 . Folding of a protein into its biologically active state is a sequen-
tial, cooperative process driven primarily by the hydrophobic ef-
fect. Folding can be assisted by chaperones.

13 . Collagen is the major fibrous protein of connective tissues. The
three left-handed helical chains of collagen form a right-handed
supercoil.

14. The compact, folded structures of proteins allow them to selectively
bind other molecules. The heme-containing proteins myoglobin
and hemoglobin bind and release oxygen. Oxygen binding to he-
moglobin is characterized by positive cooperativity and allosteric
regulation.

15 . Antibodies are multidomain proteins that bind foreign substances,
or antigens, marking them for destruction. The variable domains
at the ends of the heavy and light chains interact with the antigen.

Problems

1. Examine the following tripeptide:

H 3 N

r 3 h

^o©

(a) Label the a-carbon atoms and draw boxes around the atoms
of each peptide group.

(b) What do the R groups represent?

(d) Assuming that the chemical structure represents the correct
conformation of the peptide linkage, are the peptide groups
in the cis or the trans conformation?

(e) Which bonds allow rotation of peptide groups with respect
to each other?

2. (a) Characterize the hydrogen-bonding pattern of (1) an a helix

and (2) a collagen triple helix.

(b) Explain how the amino acid side chains are arranged in each
of these helices.

3. Explain why (1) glycine and (2) proline residues are not com-
monly found in a helices.

4. A synthetic 20 amino acid polypeptide named Betanova was de-
signed as a small soluble molecule that would theoretically form
stable /3-sheet structures in the absence of disulfide bonds. NMR
of Betanova in solution indicates that it does, in fact, form a
three-stranded antiparallel /3 sheet. Given the sequence of Be-
tanova below:

(a) Draw a ribbon diagram for Betanova indicating likely
residues for each hairpin turn between the /3 strands.

(b) Show the interactions that are expected to stabilize this
/3-sheet structure.

Betanova RGWS VQN GKYTNN GKTTEGR

5 . Each member of an important family of 250 different DNA-binding
proteins is composed of a dimer with a common protein motif.
This motif permits each DNA-binding protein to recognize and
bind to specific DNA sequences. What is the common protein
motif in the structure below?

6. Refer to Figure 4.21 to answer the following questions.

(a) To which of the four major domain categories does the middle
domain of pyruvate kinase (PK) belong (all a all / 3 , a//3, a + /3)?

(b) Describe any characteristic domain “fold” that is prominent
in this middle domain of PK.

7. Protein disulfide isomerase (PDI) markedly increases the rate of
correct refolding of the inactive ribonuclease form with random
disulfide bonds (Figure 4.35). Show the mechanism for the PDI-
catalyzed rearrangement of a nonnative (inactive) protein with
incorrect disulfide bonds to the native (active) protein with cor-
rect disulfide bonds.

132 CHAPTER 4 Proteins: Three-Dimensional Structure and Function

PDI

8. Myoglobin contains eight a helices, one of which has the follow-
ing sequence:

-Gln-Gly-Ala-Met-Asn-Fys-Ala-Leu-Glu-His-Phe-Arg-Fys-

Asp-Ile-Ala-Ala-

Which side chains are likely to be on the side of the helix that faces
the interior of the protein? Which are likely to be facing the aqueous
solvent? Account for the spacing of the residues facing the interior.

9. Homocysteine is an a-amino acid containing one more methylene
group in its side chain than cysteine (side chain = — CH 2 CH 2 SH).
Homocysteinuria is a genetic disease characterized by elevated
levels of homocysteine in plasma and urine, as well as skeletal de-
formities due to defects in collagen structure. Homocysteine re-
acts readily with allysine under physiological conditions. Show
this reaction and suggest how it might lead to defective cross-
linking in collagen.

10. The larval form of the parasite Schistosoma mansoni infects hu-
mans by penetrating the skin. The larva secretes enzymes that
catalyze the cleavage of peptide bonds between residues X and Y
in the sequence -Gly-Pro-X-Y- (X and Y can be any of several
amino acids). Why is this enzyme activity important for the parasite?

11 . (a) How does the reaction of carbon dioxide with water help ex-

plain the Bohr effect? Include the equation for the formation
of bicarbonate ion from C0 2 and water, and explain the ef-
fects of H® and C0 2 on hemoglobin oxygenation.

(b) Explain the physiological basis for the intravenous adminis-
tration of bicarbonate to shock victims.

12 . Fetal hemoglobin (Hb F) contains serine in place of the cationic
histidine at position 143 of the p chains of adult hemoglobin (Hb A).
Residue 143 faces the central cavity between the ft chains.

(a) Why does 2,3BPG bind more tightly to deoxy Hb A than to
deoxy Hb F?

(b) How does the decreased affinity of Hb F for 2,3BPG affect the
affinity of Hb F for 0 2 ?

(c) The P 50 for Hb F is 18 torr, and the P 50 for Hb A is 26 torr.
How do these values explain the efficient transfer of oxygen
from maternal blood to the fetus?

13 . Amino acid substitutions at the aft subunit interfaces of hemo-
globin may interfere with the R v T quaternary structural
changes that take place on oxygen binding. In the hemoglobin
variant Hb Ya kima> the R form is stabilized relative to the T form,
and P 50 =12 torr. Explain why the mutant hemoglobin is less effi-
cient than normal hemoglobin (P 50 = 26 torr) in delivering oxy-
gen to working muscle, where 0 2 may be as low as 10 to 20 torr.

14 . The spider venom from the Chilean Rose Tarantula ( Grammostola
spatulata) contains a toxin that is a 34-amino acid protein. It is
thought to be a globular protein that partitions into the lipid
membrane to exert its effect. The sequence of the protein is:

ECGKFMWKCKNSNDCCKDFVCSSRWKWCVFASPF

(a) Identify the hydrophobic and highly hydrophilic amino acids
in the protein.

(b) The protein is thought to have a hydrophobic face that interacts
with the lipid membrane. How can the hydrophobic amino
acids far apart in sequence interact to form a hydrophobic face?

[Adapted from Fee, S. and MacKinnon, R. (2004). Nature 430:
232-235.]

15 . Selenoprotein P is an unusual extracellular protein that contains
8-10 selenocysteine residues and has a high content of cysteine
and histidine residues. Selenoprotein P is found both as a plasma
protein and as a protein strongly associated with the surface
of cells. The association of selenoprotein P with cells is pro-
posed to occur through the interaction of selenoprotein P
with high-molecular-weight carbohydrate compounds classi-
fied as glycosaminoglycans. One such compound is heparin
(see structure on next page). Binding studies of selenoprotein P
to heparin were carried out under different pH conditions. The
results are shown in the graph on next page.

(a) How is the binding of selenoprotein P to heparin dependent
upon pH?

(b) Give possible structural reasons for the binding dependence.

(Hint: Use the information about which amino acids are
abundant in selenoprotein P in your answer) .

[Adapted from Arteel, G. E., Franken, S., Kappler, J., and Sies, H.
(2000). Biol Chem. 381:265-268.]

Selected Readings 133

16 . Gelatin is processed collagen that comes from the joints of ani-
mals. When gelatin is mixed with hot water, the triple helix struc-
ture unwinds and the chains separate, becoming random coils
that dissolve in the water. As the dissolved gelatin mixture cools,
the collagen forms a matrix that traps water; as a result, the mix-
ture turns into the jiggling semisolid mass that is recognizable as
Jell-O™. The directions on a box of gelatin include the following:
“Chill until slightly thickened, then add 1 to 2 cups cooked or raw
fruits or vegetables. Fresh or frozen pineapple must be cooked be-
fore adding.” If the pineapple is not cooked, the gelatin will not
set properly. Pineapple belongs to a group of plants called
Bromeliads and contains a protease called bromelain. Explain
why pineapple must be cooked before adding to gelatin.

17 . Hb Helsinki (HbH) is a hemoglobin mutant in which the lysine
residue at position 82 has been replaced with methionine. The
mutation is in the beta chain, and residue 82 is found in the central
cavity of hemoglobin. The oxygen binding curves for normal adult
hemoglobin (HbA, •) and HbH (■) at pH 7.4 in the presence of a
physiological concentration of 2,3BPG are shown in the graph.

[Adapted from Ikkala, E., Koskela, J., Pikkarainen, P., Rahiala, E.L.,
El-Hazmi, M. A., Nagai, K., Lang, A., and Lehmann, H. Acta Haematol.
(1976). 56:257-275.]

Explain why the curve for HbH is shifted from the curve for HbA.
Does this mutation stabilize the R or T state? What result does this
mutation have on oxygen affinity?

Selected Readings

General

Clothia, C., and Gough, J. (2009). Genomic and
structural aspects of protein evolution. Biochem. J.
419:15-28. doi: 10,1042/BJ20090122.

Creighton, T. E. (1993). Proteins: Structures and
Molecular Properties, 2nd ed. (New York: W. H.
Freeman), Chapters 4-7.

Fersht, A. (1998). Structure and Mechanism in Pro-
tein Structure (New York: W. H. Freeman).

Goodsell, D., and Olson, A. J. (1993). Soluble pro-
teins: size, shape, and function. Trends Biochem.

Sci. 18:65-68.

Goodsell, D. S., and Olson, A. J. (2000). Structural
symmetry and protein function. Annu. Rev. Biophys,
Biomolec. Struct. 29:105-153.

Kyte, J. (1995). Structure in Protein Chemistry
(New York: Garland) .

Protein Structure

Branden, C., and Tooze, J. (1991). Introduction to
Protein Structure 2nd ed. (New York: Garland).

Chothia, C., Hubbard, T., Brenner, S., Barns, H.,
and Murzin, A. (1997). Protein folds in the all-yS
and all-u classes. Annu. Rev. Biophys. Biomol. Struct.
26:597-627.

Edison, A. S. (2001). Linus Pauling and the planar
peptide bond. Nat. Struct. Biol. 8:201-202.

Harper, E. T., and Rose, G. D. (1993). Helix stop sig-
nals in proteins and peptides: the capping box.
Biochemistry 32:7605-7609.

Phizicky, E., and Fields, S. (1995). Protein-protein
interactions: methods for detection and analysis.
Microbiol. Rev. 59:94-123.

Rhodes, G. (1993). Crystallography Made Crystal
Clear (San Diego: Academic Press).

Richardson, J. S., and Richardson, D. C. (1989).
Principles and patterns of protein conformation. In
Prediction of Protein Structure and the Principles of
Protein Conformation, G. D. Fasman, ed. (New
York: Plenum), pp. 1-98.

Wang, Y., Liu, C., Yang, D., and Yu, H. (2010).
PinlAt encoding a peptidyl-prolyl cis/trans iso-
merase regulates flowering time in arabidopsis.
Molec. Cell. 37:112-122.

Uversky, V. N., and Dunker, A. K. (2010). Under-
standing protein non-folding. Biochim. Biophys.
Acta. 1804:1231-1264.

Protein Folding and Stability

Daggett, V., and Fersht, A. R. (2003). Is there a uni-
fying mechanism for protein folding? Trends
Biochem. Sci. 28:18-25.

Dill, K. A. Ozkan, S. B., Shell, M. S., and Weik, T. R.
(2008). The protein folding problem. Annu. Rev.
Biophys. 37:289-316.

Feldman, D. E., and Frydman, J. (2000). Protein
folding in vivo: the importance of molecular chap-
erones. Curr. Opin. Struct. Biol. 10:26-33.

Kryshtafovych, A., Fidelis, K., and Moult, J. (2009).
CASP8 results in context of previous experiments.
Proteins. 77(suppl 9):217-228.

Matthews, B. W. (1993). Structural and genetic
analysis of protein stability. Annu. Rev. Biochem.
62:139-160.

Saibil, H. R. and Ranson, N. A. (2002). The chaper-
onin folding machine. Trends Biochem. Sci.
27:627-632.

Sigler, P. B., Xu, Z., Rye, H. S., Burston, S. G., Fen-
ton, W. A., and Horwich, A. L. (1998). Structure
and function in GroEL- mediated protein folding.
Annu. Rev. Biochem. 67:581-608.

Smith, C. A. (2000). How do proteins fold?
Biochem. Ed. 28:76-79.

Specific Proteins

Ackers, G. K., Doyle, M. L., Myers, D., and Daugh-
erty, M. A. (1992). Molecular code for cooperativ-
ity in hemoglobin. Science 255:54-63.

Brittain, T. (2002). Molecular aspects of embry-
onic hemogloin function. Molec. Aspects Med.
23:293-342.

Davies, D. R., Padlan, E. A., and Sheriff, S. (1990).
Antibody-antigen complexes. Annu. Rev. Biochem.
59:439-473.

Eaton, W. A., Henry, E. R., Hofrichter, J., and Moz-
zarelli, A. (1999). Is cooperative binding by hemo-
globin really understood? Nature Struct. Biol.
6(4):351-3 57.

Kadler, K. (1994). Extracellular matrix 1:
fibril-forming collagens. Protein Profile
1:519-549.

Liu, R., and Ochman, H. (2007). Stepwise forma-
tion of the bacterial flagellar system. Proc. Natl.
Acad. Sci. (USA). 104:7116-7121.

Perutz, M. F. (1978). Hemoglobin structure and
respiratory transport. Sci. Am. 239(6):92-125.

Perutz, M. F., Wilkinson, A. J., Paoli, M., and
Dodson, G. G. (1998). The stereochemical
mechanism of the cooperative effects in
hemoglobin revisited. Annu. Rev. Biophys. Biomol.
Struct. 27:1-34.

Properties of Enzymes

W e have seen how the three-dimensional shapes of proteins allow them to
serve structural and transport roles. We now discuss their functions as en-
zymes. Enzymes are extraordinarily efficient, selective, biological catalysts.
Every living cell has hundreds of different enzymes catalyzing the reactions essential for
life — even the simplest living organisms contain hundreds of different enzymes. In
multicellular organisms, the complement of enzymes differentiates one cell type from
another but most of the enzymes we discuss in this book are among the several hundred
common to all cells. These enzymes catalyze the reactions of the central metabolic path-
ways necessary for the maintenance of life.

In the absence of the enzymes, metabolic reactions will not proceed at significant
rates under physiological conditions. The primary role of enzymes is to enhance the
rates of these reactions to make life possible. Enzyme -catalyzed reactions are 10 3 to 10 20
times faster than the corresponding uncatalyzed reactions. A catalyst is defined as a
substance that speeds up the attainment of equilibrium. It may be temporarily changed
during the reaction but it is unchanged in the overall process since it recycles to partici-
pate in multiple reactions. Reactants bind to a catalyst and products dissociate from it.
Note that a catalyst does not change the position of the reactions equilibrium (i.e., it
does not make an unfavorable reaction favorable). Instead, it lowers the amount of en-
ergy needed in order for the reaction to proceed. Catalysts speed up both the forward
and reverse reactions by converting a one- or two-step process into several smaller steps
each needing less energy than the uncatalyzed reaction.

Enzymes are highly specific for the reactants, or substrates, they act on, but the de-
gree of substrate specificity varies. Some enzymes act on a group of related substrates,
and others on only a single compound. Many enzymes exhibit stereospecificity meaning

I was awed by enzymes and fell
instantly in love with them. I have
since had love affairs with many
enzymes (none as enduring as with
DNA polymerase ), but I have never
met a dull or disappointing one.

—Arthur Kornberg (2001)

KEY CONCEPT

Catalysts speed up the rate of
forward and reverse reactions but
they don’t change the equilibrium
concentrations.

Top:The enzyme acetylcholinesterase with the reversible inhibitor donepezil hydrochloride (Aricept; shown in red) occupy-
ing the active site. Aricept is used to improve mental functioning in patients with Alzheimer’s disease. It is thought to act
by inhibiting the breakdown of the neurotransmitter acetylcholine in the brain, thus prolonging the neurotransmitter ef-
fects. (It does not, however, affect the course of the disease.) [PDB 1EVE]

134

Properties of Enzymes

135

▲ Enzyme reaction. This is a large-scale enzyme reaction where milk is being curdled to make
Appenzeller cheese. The reaction is catalyzed by rennet (rennin), which was originally derived from
cow stomach. Rennet contains the enzyme chymosin, a protease that cleaves the milk protein
casein between phenylalanine and methionine residues. The reaction releases a hydrophobic
fragment of casein that aggregates and precipitates forming curd.

that they act on only a single stereoisomer of the substrate. Perhaps the most important
aspect of enzyme specificity is reaction specificity — that is, the lack of formation of
wasteful by-products. Reaction specificity is reflected in the exceptional purity of prod-
uct (essentially 100%) — much higher than the purity of products of typical catalyzed
reactions in organic chemistry. The specificity of enzymes not only saves energy for cells
but also precludes the buildup of potentially toxic metabolic by-products.

Enzymes can do more than simply increase the rate of a single, highly specific reac-
tion. Some can also combine, or couple, two reactions that would normally occur sepa-
rately. This property allows the energy gained from one reaction to be used in a second
reaction. Coupled reactions are a common feature of many enzymes — the hydrolysis of
ATP, for example, is often coupled to less favorable metabolic reactions.

Some enzymatic reactions function as control points in metabolism. As we will see,
metabolism is regulated in a variety of ways including alterations in the concentrations
of enzymes, substrates, and enzyme inhibitors and modulation of the activity levels of
certain enzymes. Enzymes whose activity is regulated generally have a more complex
structure than unregulated enzymes. With few exceptions, regulated enzymes are
oligomeric molecules that have separate binding sites for substrates and effectors, the
compounds that act as regulatory signals. The fact that enzyme activity can be regulated
is an important property that distinguishes biological catalysts from those encountered
in a chemistry lab.

The word enzyme is derived from a Greek word meaning “in yeast.” It indicates that
these catalysts are present inside cells. In the late 1800s, scientists studied the fermentation
of sugars by yeast cells. Vitalists (who maintained that organic compounds could be
made only by living cells) said that intact cells were needed for fermentation. Mechanists
claimed that enzymes in yeast cells catalyze the reactions of fermentation. The latter
conclusion was supported by the observation that cell- free extracts of yeast can catalyze
fermentation. This finding was soon followed by the identification of individual reactions
and the enzymes that catalyze them.

A generation later, in 1926, James B. Sumner crystallized the first enzyme (urease)
and proved that it is a protein. Five more enzymes were purified in the next decade and
also found to be proteins: pepsin, trypsin, chymotrypsin, carboxypeptidase, and Old
Yellow Enzyme (a flavoprotein NADPH oxidase). Since then, almost all enzymes have
been shown to be proteins or proteins plus cofactors. Certain RNA molecules also ex-
hibit catalytic activity but they are not usually referred to as enzymes.

Some of the first biochemistry depart-
ments in universities were called
Departments of Zymology.

Catalytic RNA molecules are discussed
in Chapters 21 and 22.

136 CHAPTER 5 Properties of Enzymes

▲ Crystals of a bacterial ( Shewanella
oneidensis ) homologue of Old Yellow Enzyme.

(Courtesy of J. Elegheert and S. N.
Savvides)

We begin this chapter with a description of enzyme classification and nomencla-
ture. Next, we discuss kinetic analysis (measurements of reaction rates) emphasizing
how kinetic experiments can reveal the properties of an enzyme and the nature of the
complexes it forms with substrates and inhibitors. Finally, we describe the principles of
inhibition and activation of regulatory enzymes. Chapter 6 explains how enzymes work
at the chemical level and uses serine proteases to illustrate the relationship between pro-
tein structure and enzymatic function. Chapter 7 is devoted to the biochemistry of
coenzymes, the organic molecules that assist some enzymes in their catalytic roles by
providing reactive groups not found on amino acid side chains. In the remaining chapters
we will present many other examples illustrating the four main properties of enzymes:
(1) they function as catalysts, (2) they catalyze highly specific reactions, (3) they can
couple reactions, and (4) their activity can be regulated.

5.1 The Six Classes of Enzymes

Most of the classical metabolic enzymes are named by adding the suffix -ase to the
name of their substrates or to a descriptive term for the reactions they catalyze. For ex-
ample, urease has urea as a substrate. Alcohol dehydrogenase catalyzes the removal of
hydrogen from alcohols (i.e., the oxidation of alcohols). A few enzymes, such as trypsin
and amylase, are known by their historic names. Many newly discovered enzymes are
named after their genes or for some nondescriptive characteristic. For example, RecA is
named after the recA gene and HSP70 is a heat shock protein — both enzymes catalyze
the hydrolysis of ATR

A committee of the International Union of Biochemistry and Molecular Biology
(IUBMB) maintains a classification scheme that categorizes enzymes according to the
general class of organic chemical reaction that is catalyzed. The six categories —
oxidoreductases, transferases, hydrolases, lyases, isomerases, and ligases — are defined
below with an example of each type of enzyme. The IUBMB classification scheme as-
signs a unique number, called the enzyme classification number, or EC number, to
each enzyme. IUBMB also assigns a unique systematic name to each enzyme; it may be
different from the common name of an enzyme. This book usually refers to enzymes
by their common names.

1. Oxidoreductases catalyze oxidation-reduction reactions. Most of these enzymes are
commonly referred to as dehydrogenases. Other enzymes in this class are called oxi-
dases, peroxidases, oxygenases, or reductases. There is a trend in biochemistry to
refer to more and more of these enzymes by their systematic name, oxidoreduc-
tases, rather than the more common names in the older biochemical literature.
One example of an oxidoreductase is lactate dehydrogenase (EC 1.1.1.27) also
called lactate:NAD oxidoreductase. This enzyme catalyzes the reversible conversion
of L-lactate to pyruvate. The oxidation of L-lactate is coupled to the reduction of
the coenzyme nicotinamide adenine dinucleotide (NAD®).

COO

HO — C — H + NAD

ch 3

L-Lactate

Lactate

dehydrogenase

COO

C = 0 + NADH + H

(5.1)

CH 3

Pyruvate

2. Transferases catalyze group transfer reactions and many require the presence of
coenzymes. In group transfer reactions a portion of the substrate molecule usually
binds covalently to the enzyme or its coenzyme. This group includes kinases,
enzymes that catalyze the transfer of a phosphoryl group from ATP. Alanine
transaminase, whose systematic name is L- alanine: 2 -oxyglutarate aminotransferase

5.1 The Six Classes of Enzymes 137

BOX 5.1 ENZYME CLASSIFICATION NUMBERS

The enzyme classification number for malate dehydrogenase
is EC 1.1.1.37. This enzyme has an activity similar to that of
lactate dehydrogenase described under oxidoreductases (see
Figure 4.23, Box 13.3).

The first number identifies this enzyme as a member of the
first class of enzymes (oxidoreductases). The second number
identifies the substrate group that malate dehydrogenase recog-
nizes. Subclass 1.1 means that the substrate is a HC — OH
group. The third number specifies the electron acceptor for
this class of enzymes. Subclass 1.1.1 is for enzymes that use
NAD + or NADP + as an acceptor. The final number means that
malate dehydrogenase is the 37th enzyme in this category.

Compare the EC number
of malate dehydrogenase with
that of lactate dehydrogenase to
see how similar enzymes have
similar classification numbers.

Accurate enzyme identifi-
cation and classification is an
important and essential part of
modern biological databases.
The entire classification data-
base can be seen at www.chem.
qmul.ac.uk/iubmb/enzyme/.

(EC 2.6. 1.2), is a typical transferase. It transfers an amino group from L-alanine to
a-ketoglutarate (2-oxoglutarate) .

COO w COO

i i

-c — H + C=0

Alanine transaminase

< =±

coo°

I ©

C = 0 + H,N-

,©

ch 3

L-Alanine

(CH 2 ) 2

COO

CH 3

Pyruvate

COO

-c — H

(CH 2 ) 2

(5.2)

u-Ketoglutarate

coo°

L-Glutamate

3. Hydrolases catalyze hydrolysis. They are a special class of transferases with water
serving as the acceptor of the group transferred. Pyrophosphatase is a simple exam-
ple of a hydrolase. The systematic name of this enzyme is diphosphate phosphohy-
drolase (EC 3. 6. 1.1).

0 .

O-

-o— p— o + h 2 o

L©

Pyrophosphatase

°o o'-

Pyrophosphate

2 HO — P — O

o®

Phosphate

(5.3)

4. Lyases catalyze lysis of a substrate generating a double bond in nonhydrolytic,
nonoxidative, elimination reactions. In the reverse direction, lyases catalyze the ad-
dition of one substrate to the double bond of a second substrate. Pyruvate decar-
boxylase belongs to this class of enzymes since it splits pyruvate into acetaldehyde
and carbon dioxide. The systematic name for pyruvate decarboxylase, 2-oxo-acid
carboxy-lyase (EC 4. 1.1.1), is rarely used.

C = 0 + H

CH 3

Pyruvate

Pyruvate H O

decarboxylase \ #

> C +

CH 3

Acetaldehyde

0 = C=0

Carbon

dioxide

(5.4)

5. Isomerases catalyze structural change within a single molecule (isomerization reac-
tions). Because these reactions have only one substrate and one product, they are
among the simplest enzymatic reactions. Alanine racemase (EC 5. 1.1.1) is an

▲ Distribution of all known enzymes by EC
classification number. 1. oxidoreductases;
2. transferases; 3. hydrolases; 4. lyases;
5. isomerases; 6. ligases.

138 CHAPTER 5 Properties of Enzymes

isomerase that catalyzes the interconversion of L-alanine and D-alanine. The com-
mon name is the same as the systematic name.

coo 0

H 3 N — C — H

ch 3

L-Alanine

Alanine

racemase

coo 0

I ©

H — C — NH 3

ch 3

D-Alanine

(5.5)

6. Ligases catalyze ligation, or joining, of two substrates. These reactions require the
input of chemical potential energy in the form of a nucleoside triphosphate
such as ATP. Ligases are usually referred to as synthetases. Glutamine synthetase, or
L- glutamate: ammonia ligase (ADP-forming) (EC 6.3. 1.2), uses the energy of ATP
hydrolysis to join glutamate and ammonia to produce glutamine.

The human genome contains genes for
about 1000 different enzymes catalyz-
ing reactions in several hundred meta-
bolic pathways (humancyc.org/). Since
many enzymes have multiple subunits
there are about 3000 different genes
devoted to making enzymes. We have
about 20,000 genes so most of the
genes in our genome do not encode
enzymes or enzyme subunits.

coo 0

H 3 N — C — H

I + ATP + NH 4 @
(CH 2 ) 2

Glutamine

synthetase

coo°

H 3 N — C — H

I + ADP + P|

(CH 2 ) 2 (5.6)

->©

L-Glutamate

S \

O NH
L-Glutamine

From the examples given above we see that most enzymes have more than one sub-
strate although the second substrate may be only a molecule of water or a proton. Al-
though enzymes catalyze both forward and reverse reactions, one-way arrows are often
used when the equilibrium favors a great excess of product over substrate. Remember
that when a reaction reaches equilibrium the enzyme must be catalyzing both the for-
ward and reverse reactions at the same rate.

Recall that concentrations are indicat-
ed by square brackets: [P] signifies the
concentration of product, [E] the con-
centration of enzyme, and [S] the con-
centration of the substrate.

5.2 Kinetic Experiments Reveal Enzyme Properties

We begin our study of enzyme properties by examining the rates of enzyme -catalyzed
reactions. Such studies fall under the category of enzyme kinetics (from the Greek
kinetikos , “moving”). This is an appropriate place to begin since the most important
property of enzymes is that they act as catalysts, speeding up the rates of reactions. En-
zyme kinetics provides indirect information about the specificities and catalytic mecha-
nisms of enzymes. Kinetic experiments also reveal whether an enzyme is regulated.

Most enzyme research in the first half of the 20th century was limited to kinetic ex-
periments. This research revealed how the rates of reactions are affected by variations in
experimental conditions or changes in the concentration of enzyme or substrate. Before
discussing enzyme kinetics in depth, let’s review the principles of kinetics for
nonenzymatic chemical systems. These principles are then applied to enzymatic reactions.

A. Chemical Kinetics

Kinetic experiments examine the relationship between the amount of product (P)
formed in a unit of time (A[P]/At) and the experimental conditions under which the re-
action takes place. The basis of most kinetic measurements is the observation that the
rate, or velocity (v), of a reaction varies directly with the concentration of each reactant
(Section 1.4). This observation is expressed in a rate equation. For example, the rate
equation for the nonenzymatic conversion of substrate (S) to product in an isomeriza-
tion reaction is written as

A[P]

v = k[S]

(5.7)

5.2 Kinetic Experiments Reveal Enzyme Properties 139

The rate equation reflects the fact that the velocity of a reaction depends on the concen-
tration of the substrate ([S]). The symbol k is the rate constant and indicates the speed
or efficiency of a reaction. Each reaction has a different rate constant. The units of the
rate constant for a simple reaction are s _1 .

As a reaction proceeds, the amount of product ([P]) increases and the amount of
substrate ([S]) decreases. An example of the progress of several reactions is shown in
Figure 5.1a. The velocity is the slope of the progress curve over a particular interval of time.
The shape of the curves indicates that the velocity is decreasing over time as expected
since the substrate is being depleted.

In this hypothetical example, the velocity of the reaction might eventually become
zero when the substrate is used up. This would explain why the curve flattens out at ex-
tended time points. (See below for another explanation.) We are interested in the rela-
tionship between substrate concentration and the velocity of a reaction since if we
know these two values we can use Equation 5.7 to calculate the rate constant. The only
accurate substrate concentration is the one we prepare at the beginning of the experi-
ment because the concentration changes during the experiment. The velocity of the re-
action at the very beginning is the value that we want to know. This value represents the
rate of the reaction at a known substrate concentration before it changes.

The initial velocity (v 0 ) can be determined from the slope of the progress curves
(Figure 5.1a) or from the derivatives of the curves. A graph of initial velocity versus sub-
strate concentration at the beginning of the experiment gives a straight line as shown in
Figure 5.1b. The slope of the curve in Figure 5.1b is the rate constant.

The experiment shown in Figure 5.1 will only determine the forward rate constant
since the data were collected under conditions where there was no reverse reaction. This
is another important reason for calculating initial velocity (v 0 ) rather than the rate at
later time points. In a reversible reaction, the flattening of the progress curves does not
represent zero velocity. Instead, it simply indicates that there is no net increase in prod-
uct over time because the reaction has reached equilibrium.

A better description of our simple reaction would be

S P (5.8)

/c_i

For a more complicated single-step reaction, such as the reaction S x + S 2 — » Pi + P 2 > the
rate is determined by the concentrations of both substrates. If both substrates are pres-
ent at similar concentrations, the rate equation is

v= /c[S n ][S 2 ] (5.9)

The rate constant for reactions involving two substrates has the units M -1 s -1 . These
rate constants can be easily determined by setting up conditions where the concentra-
tion of one substrate is very high and the other is varied. The rate of the reaction will
depend on the concentration of the rate-limiting substrate.

B. Enzyme Kinetics

One of the first great advances in biochemistry was the discovery that enzymes bind
substrates transiently. In 1894, Emil Fischer proposed that an enzyme is a rigid tem-
plate, or lock, and that the substrate is a matching key. Only specific substrates can fit
into a given enzyme. Early studies of enzyme kinetics confirmed that an enzyme (E)
binds a substrate to form an enzyme-substrate complex (ES). ES complexes are formed
when ligands bind noncovalently in their proper places in the active site. The substrate
interacts transiently with the protein catalyst (and with other substrates in a multisub-
strate reaction) on its way to forming the product of the reaction.

Lets consider a simple enzymatic reaction; namely, the conversion of a single sub-
strate to a product. Although most enzymatic reactions have two or more substrates, the
general principles of enzyme kinetics can be described by assuming the simple case of
one substrate and one product.

E + S > ES > E + P (5.10)

0.05 M 0.1 M 0.2 M

[S]

▲ Figure 5.1

Rate of a simple chemical reaction, (a) The

amount of product produced over time is
plotted for several different initial substrate
concentrations. The initial velocity i/ 0 is the
slope of the progress curve at the beginning
of the reaction, (b) The initial velocity as a
function of initial substrate concentration.
The slope of the curve is the rate constant.

KEY CONCEPT

The rate or velocity of a reaction depends
on the concentration of substrate.

140 CHAPTER 5 Properties of Enzymes

KEY CONCEPT

The enzyme-substrate complex (ES) is a
transient intermediate in an enzyme
catalyzed reaction.

[E]

▲ Figure 5.2

Effect of enzyme concentration ([E]), on the
initial velocity (v) of an enzyme-catalyzed
reaction at a fixed, saturating [S]. The

reaction rate is affected by the concentra-
tion of enzyme but not by the concentration
of the other reactant, S.

Time (t)

▲ Figure 5.3 Progress curve for an enzyme-
catalyzed reaction. [P], the concentration of
product, increases as the reaction proceeds.
The initial velocity of the reaction, i/ 0 , is the
slope of the initial linear portion of the
curve. Note that the rate of the reaction
doubles when twice as much enzyme
(2E, upper curve) is added to an otherwise
identical reaction mixture.

This reaction takes place in two distinct steps — the formation of the enzyme-substrate
complex and the actual chemical reaction accompanied by the dissociation of the en-
zyme and product. Each step has a characteristic rate. The overall rate of an enzymatic
reaction depends on the concentrations of both the substrate and the catalyst (enzyme).
When the amount of enzyme is much less than the amount of substrate the reaction
will depend on the amount of enzyme.

The straight line in Figure 5.2 illustrates the effect of enzyme concentration on the
reaction velocity in a pseudo first-order reaction. The more enzyme present, the faster
the reaction. These conditions are used in enzyme assays to determine the concentra-
tions of enzymes. The concentration of enzyme in a test sample can be easily deter-
mined by comparing its activity to a reference curve similar to the model curve in
Figure 5.2. Under these experimental conditions, there are sufficient numbers of sub-
strate molecules so that every enzyme molecule binds a molecule of substrate to form
an ES complex, a condition called saturation of E with S. Enzyme assays measure the
amount of product formed in a given time period. In some assay methods, a recording
spectrophotometer can be used to record data continuously; in other methods, samples
are removed and analyzed at intervals. The assay is performed at a constant pH and
temperature, generally chosen for optimal enzyme activity or for approximation to
physiological conditions.

If we begin an enzyme-catalyzed reaction by mixing substrate and enzyme then
there is no product present during the initial stages of the reaction. Under these condi-
tions we can ignore the reverse reaction where P binds to E and is converted to S. The
reaction can be described by

k-\ k?

E + S ES — ^ E + P (5.11)

/C_!

The rate constants k\ and k- X in Reaction 5.1 1 govern the rates of association of S with E
and dissociation of S from ES, respectively. This first step is an equilibrium binding in-
teraction similar to the binding of oxygen to hemoglobin. The rate constant for the sec-
ond step is k 2 , the rate of formation of product from ES. Note that conversion of the ES
complex to free enzyme and product is shown by a one-way arrow because the rate of
the reverse reaction (E + P — » EP) is negligible at the start of a reaction. The velocity
measured during this short period is the initial velocity (v 0 ) described in the previous
section. The formation and dissociation of ES complexes are usually very rapid reac-
tions because only noncovalent bonds are being formed and broken. In contrast, the
conversion of substrate to product is usually rate limiting. It is during this step that the
substrate is chemically altered.

Enzyme kinetics differs from simple chemical kinetics because the rates of enzyme-
catalyzed reactions depend on the concentration of enzyme and the enzyme is neither a
product nor a substrate of the reaction. The rates also differ because substrate has to
bind to enzyme before it can be converted to product. In an enzyme -catalyzed reaction,
the initial velocities are obtained from progress curves, just as they are in chemical reac-
tions. Figure 5.3 shows the progress curves at two different enzyme concentrations in
the presence of a high initial concentration of substrate ([S] » [E] ). In this case, the
rate of product formation depends on enzyme concentration and not on the substrate
concentration. Data from experiments such as those shown in Figure 5.3 can be used to
plot the curve shown in Figure 5.2.

5.3 The Michaelis-Menten Equation

Enzyme- catalyzed reactions, like any chemical reaction, can be described mathemati-
cally by rate equations. Several constants in the equations indicate the efficiency and
specificity of an enzyme and are therefore useful for comparing the activities of several
enzymes or for assessing the physiological importance of a given enzyme. The first rate
equations were derived in the early 1900s by examining the effects of variations in sub-
strate concentration. Figure 5.4 a shows a typical result where the initial velocity (v 0 ) of
a reaction is plotted against the substrate concentration ( [S] ).

5.3 The Michael is-Menten Equation 141

The data can be explained by the reaction shown in Reaction 5.1 1. The first step is a
bimolecular interaction between the enzyme and substrate to form an ES complex. At
high substrate concentrations (right-hand side of the curve in Figure 5.4) the initial ve-
locity doesn’t change very much as more S is added. This indicates that the amount of
enzyme has become rate-limiting in the reaction. The concentration of enzyme is an
important component of the overall reaction as expected for formation of an ES
complex. At low substrate concentrations (left-hand side of the curve in Figure 5.4), the
initial velocity is very sensitive to changes in the substrate concentration. Under these
conditions most enzyme molecules have not yet bound substrate and the formation of
the ES complex depends on the substrate concentration.

The shape of the v 0 vs. [S] curve is that of a rectangular hyperbola. Hyperbolic
curves indicate processes involving simple dissociation as we saw for the dissociation of
oxygen from oxymyoglobin (Section 4.13B). This is further evidence that the simple re-
action under study is bimolecular involving the association of E and S to form an ES
complex. The equation for a rectangular hyperbola is

y = VT~ x

(5.12)

where a is the asymptote of the curve (the value of y at an infinite value of x) and b is
the point on the x axis corresponding to a value of a/2. In enzyme kinetic experiments,
y - v 0 and x = [S]. The asymptote value (a) is called l/ max . It’s the maximum velocity of
the reaction at infinitely large substrate concentrations. We often show the V max value
on v 0 vs. [S] plots but if you look at the figure it’s not obvious why this particular as-
ymptote was chosen. One of the characteristics of hyperbolic curves is that the curve
seems to flatten out at moderate substrate concentrations at a level that seems far less
than the V^ax value. The true Vm ax is n °t determined by trying to estimate the position
of the asymptote from the shape of the curve; instead, it is precisely and correctly deter-
mined by fitting the data to the general equation for a rectangular hyperbola.

The b term in the general equation for a rectangular hyperbola is called the
Michaelis constant (K m ) defined as the concentration of substrate when v 0 is equal to
one -half Vm ax (Figure 5.4b). The complete rate equation is

Knax[S]

/C m + [S]

(5.13)

This is called the Michaelis-Menten equation, named after Leonor Michaelis and Maud
Menten. Note how the general form of the equation compares to Equation 5.12. The
Michaelis-Menten equation describes the relationship between the initial velocity of a
reaction and the substrate concentration. In the following section we derive the
Michaelis-Menten equation by a kinetic approach and then consider the meaning of
the various constants.

A. Derivation of the Michaelis-Menten Equation

One common derivation of the Michaelis-Menten equation is termed the steady state
derivation. It was proposed by George E. Briggs and J. B. S. Haldane. This derivation
postulates a period of time (called the steady state) during which the ES complex is
formed at the same rate that it decomposes so that the concentration of ES is constant.
The initial velocity is used in the steady state derivation because we assume that the
concentration of product ( [P] ) is negligible. The steady state is a common condition for
metabolic reactions in cells.

If we assume a constant steady state concentration of ES then the rate of formation
of product depends on the rate of the chemical reaction and the rate of dissociation of P
from the enzyme. The rate limiting step is the right-hand side of Reaction 5.11 and the
velocity depends on the rate constant k 2 and the concentration of ES.

ES — E + P v 0 = k 2 [ES] (5.14)

(a)

0 [Si

(b)

▲ Figure 5.4

Plots of initial velocity (v 0 ) versus substrate
concentration ([S]) for an enzyme-catalyzed
reaction, (a) Each experimental point is
obtained from a separate progress curve
using the same concentration of enzyme.
The shape of the curve is hyperbolic. At
low substrate concentrations, the curve ap-
proximates a straight line that rises steeply.
In this region of the curve, the reaction is
highly dependent on the concentration of
substrate. At high concentrations of sub-
strate, the enzyme is almost saturated, and
the initial rate of the reaction does not
change much when substrate concentration
is further increased, (b) The concentration
of substrate that corresponds to half-maxi-
mum velocity is called the Michaelis con-
stant (K m ). The enzyme is half-saturated
when S = K m .

142 CHAPTER 5 Properties of Enzymes

▲ Leonor Michaelis (1875-1949).

The steady-state derivation solves Equation 5.14 for [ES] using terms that can be meas-
ured such as the rate constant, the total enzyme concentration ([E] tota i), and the sub-
strate concentration ([S]). [S] is assumed to be greater than [E] tota i but not necessarily
saturating. For example, soon after a small amount of enzyme is mixed with substrate [ES]
becomes constant because the overall rate of decomposition of ES (the sum of the rates
of conversion of ES to E + S and to E + P) is equal to the rate of formation of the ES
complex from E + S. The rate of formation of ES from E + S depends on the concentra-
tion of free enzyme (enzyme molecules not in the form of ES) which is [E] tota i — [ES].
The concentration of the ES complex remains constant until consumption of S causes
[S] to approach [E] tota p We can express these statements as a mathematical equation.

Rate of ES formation = Rate of ES decomposition

*l([E]total - [ES])[S] = (*_, + * 2 )[ES]

Equation 5.15 is rearranged to collect the rate constants.

k-i+k 2 _ _ l[E]totai - [ES]2[S]

_ ki " m ” [ES f

(5.15)

(5.16)

The ratio of rate constants on the left-hand side of Equation 5.16 is the Michaelis con-
stant, K m . Next, this equation is solved for [ES] in several steps.

[ES ]K m = ([E] tota | - [ES])[S] (5.17)

Expanding,

[ES]K m = ([E] tota |[S]) - ([ES][S]) (5.18)

Collecting [ES] terms,

and

[ES](K m + [S]) = [E] tota |[S]

[E]total[S]
K m + [S]

(5.19)

(5.20)

▼ Maud Menten (1879-1960).

I MAUD LEONORA MENTEN 1879-1960

J An outstanding medical scientist. Maud Menten was born in
Port Lamb ton. She graduated in medicine from the University
of Toronto in 1907 and four years later he came one of (he first
Canadian women to receive a medical doctorate. In 19T5 in

! Germany collaboration with Leonor Michaelis on the' behaviour
ot enzymes resulted in the Michaelis -Menten equation, a basic
biochemical concept which brought them international rccog-

! nition. Menten continued her brilliant career as a pathologist
at the University of Pittsburgh from 19 18* publishing exten-
sively on medical and biochemical subjects. Her many achieve-
ments included important co-discoveries relating to blood sugar,
I haemoglobin, and kidney functions. Between 1951 pud 1954
I she conducted cancer research in British Columbia and re-
I turned to Ontario six years before she died.

br .Sp 0**19 H., Twxfafc* Kitirtu =1 C*l«w i*t O'**"*

5.3 The Michael is-Menten Equation

143

Equation 5.20 describes the steady-state ES concentration using terms that can be
measured in an experiment. Substituting the value of [ES] into the velocity equation
(Equation 5.14) gives

V'o = MES] =

fc2[E]total[S]

K m + [S]

(5.21)

As indicated by Figure 5.4a, when the concentration of S is very high the enzyme is
saturated and essentially all the molecules of E are present as ES. Adding more S has al-
most no effect on the reaction velocity. The only way to increase the velocity is to add
more enzyme. Under these conditions the velocity is at its maximum rate (Umax) and
this velocity is determined by the total enzyme concentration and the rate constant k 2 .
Thus, by definition,

Knax ^2[E]total

(5.22)

Substituting this in Equation 5.21 gives the most familiar form of the
Michaelis-Menten equation.

^0 =

US]

/C m + [S]

(5.23)

KEY CONCEPT

The constant /r cat is the number of moles
of substrate converted to product per
second per mole of enzyme.

We’ve already seen that this form of the Michaelis-Menten equation adequately de-
scribes the data from kinetic experiments. In this section we’ve shown that the same
equation can be derived from a theoretical consideration of the implications of Reac-
tion 5.11, the equation for an enzyme -catalyzed reaction. The agreement between the-
ory and data gives us confidence that the theoretical basis of enzyme kinetics is sound.

B. The Catalytic Constant /r cat

At high substrate concentration, the overall velocity of the reaction is V max and the rate
is determined by the enzyme concentration. The rate constant observed under these
conditions is called the catalytic constant, /r cat , defined as

Knax = ^cat[E]total ^cat = ^ ~ (5.24)

where fc cat represents the number of moles of substrate converted to product per second
per mole of enzyme (or per mole of active site for a multisubunit enzyme) under satu-
rating conditions. In other words, fc cat indicates the maximum number of substrate
molecules converted to product each second by each active site. This is often called
the turnover number. The catalytic constant measures how quickly a given enzyme can
catalyze a specific reaction — it’s a very useful way of describing the effectiveness of
an enzyme. The unit for fc cat is s _1 and the reciprocal of fc cat is the time required for
one catalytic event. Note that the enzyme concentration must be known in order to
calculate fc cat .

For a simple reaction, such as Reaction 5.1 1, the rate-limiting step is the conversion
of substrate to product and the dissociation of product from the enzyme (ES — > E + P).
Under these conditions fc cat is equal to k 2 (Equation 5.14). Many enzyme reactions are
more complex. If one step is clearly rate-limiting then its rate constant is the fc cat for that
reaction. If the mechanism is more complex then fc cat may be a combination of several
different rate constants. This is why we need a different rate constant (fc cat ) to describe
the overall rate of the enzyme -catalyzed reaction. In most cases you can assume that fc cat
is a good approximation of k 2 .

Representative values of fc cat are listed in Table 5.1. Most enzymes are potent catalysts
with fc cat values of 10 2 to 10 3 s _1 . This means that at high substrate concentrations a single

Table 5.1 Examples of catalytic constants

Enzyme

*cat(s V

Papain

Ribonuclease

10 2

Carboxypeptidase

10 2

Trypsin

10 2 (to 10 3 )

Acetylcholinesterase

10 3

Kinases

10 3

Dehydrogenases

10 3

Transaminases

10 3

Carbonic anhydrase

10 6

Superoxide dismutase

10 6

Catalase

10 7

*The catalytic constants are given only as orders
of magnitude.

144 CHAPTER 5 Properties of Enzymes

▲ Substrate binding. Pyruvate carboxylase
binds pyruvate, HC0 3 “ and ATP. The
structure of the active site of the yeast
( Saccharomyces cerevisiae) enzyme is
shown here with a bound molecule of
pyruvate (space-filling representation) and
the cofactor biotin (bal l-and-stick). The K m
value for pyruvate binding is 4 x 1CT 4 M.
The K m values for HC 03 ~ and ATP binding
are 1 x 1CT 3 M and 6 x 1CT 5 M.

[PDB 2VK1]

enzyme molecule will convert 100-1000 molecules of substrate to product every second.
This rate is limited by a number of factors that will be discussed in the next chapter
(Chapter 6: Mechanisms of Enzymes).

Some enzymes are extremely rapid catalysts with k cat values of 10 6 s _1 or greater.
Mammalian carbonic anhydrase, for example, must act very rapidly in order to main-
tain equilibrium between aqueous C0 2 and bicarbonate (Section 2.10). As we will see in
Section 6.4B, superoxide dismutase and catalase are responsible for rapid decomposi-
tion of the toxic oxygen metabolites superoxide anion and hydrogen peroxide, respec-
tively. Enzymes that catalyze a million reactions per second often act on small substrate
molecules that diffuse rapidly inside the cell.

C. The Meanings of K m

The Michaelis constant has a number of meanings. Equation 5.16 defined K m as the
ratio of the combined rate constants for the breakdown of ES divided by the constant
for its formation. If the rate constant for product formation ( k 2 ) is much smaller than
either k x or k- X , as is often the case, k 2 can be neglected and K m is equivalent to k-i/k x .
In this case K m is the same as the equilibrium constant for dissociation of the ES com-
plex to E +S. Thus, K m becomes a measure of the affinity of E for S. The lower the value
of K m , the more tightly the substrate is bound. K m is also one of the parameters that
determines the shape of the v 0 vs. [S] curve shown in Figure 5.4b. It is the substrate con-
centration when the initial velocity is one-half the V max value. This meaning follows
directly from the general equation for a rectangular hyperbola.

K m values are sometimes used to distinguish between different enzymes that cat-
alyze the same reaction. For example, mammals have several different forms of lactate
dehydrogenase, each with a distinct K m value. Although it is useful to think of K m
as representing the equilibrium dissociation constant for ES, this is not always valid.
For many enzymes K m is a more complex function of the rate constants. This is espe-
cially true when the reaction occurs in more than two steps.

Typical K m values for enzymes range from 10 -2 to 10 -5 M. Since these values often
represent apparent dissociation constants their reciprocal is an apparent association
(binding) constant. You can see by comparison with protein-protein interactions
(Section 4.9) that the binding of enzymes to substrates is much weaker.

KEY CONCEPT

K m is the substrate concentration when
the rate of the reaction is one-half the
I/max value. It is often an approximation of
the equilibrium dissociation constant of
the reaction ES E + S.

5.4 Kinetic Constants Indicate Enzyme Activity
and Catalytic Proficiency

We’ve seen that the kinetic constants K m and k CdLt can be used to gauge the relative activ-
ities of enzymes and substrates. In most cases, K m is a measure of the stability of the ES
complex and k Q2X is similar to the rate constant for the conversion of ES to E + P when
the substrate is not limiting (region A in Figure 5.5). Recall that k cat is a measure of the
catalytic activity of an enzyme indicating how many reactions a molecule of enzyme
can catalyze per second.

Examine region B of the hyperbolic curve in Figure 5.5. The concentration of S is
very low and the curve approximates a straight line. Under these conditions, the reac-
tion rate depends on the concentrations of both substrate and enzyme. In chemical
terms, this is a second-order reaction and the velocity depends on a second-order rate
constant defined by

v 0 = *[E][S] (5.25)

We are interested in knowing how to determine this second- order rate constant since it
tells us the rate of the enzyme -catalyzed reaction under physiological conditions. When
Michaelis and Menten first wrote the full rate equation they used the form that included
k cat [E\ total rather than U max (Equation 5.24). Now that we understand the meaning of /c cat

5.5 Measurement of K m and k max 145

▲ Figure 5.5 Meanings of /r cat and k ca ^/K m . The catalytic constant (/r cat ) is the rate constant for con-
version of the ES complex to E + P. It is measured most easily when the enzyme is saturated with
substrate (region A on the Michael is-Menten curve shown). The ratio k cat /K m is the rate constant
for the conversion of E + S to E + P at very low concentrations of substrate (region B). The reac-
tions measured by these rate constants are summarized below the graph.

we can substitute fc cat [E] total i n the Michaelis-Menten equation (Equation 5.23) in place
of V max . If we consider only the region of the Michaelis-Menten curve at a very low [S]
then this equation can be simplified by neglecting the [S] in the denominator since [S]
is much less than K m .

WE][S]
+ [S]

f*[E][S]

(5.26)

Comparing Equations 5.25 and 5.26 reveals that the second-order rate constant is
closely approximated by k cat /K m . Thus, the ratio k cat /K m is an apparent second-order
rate constant for the formation of E + P from E + S when the overall reaction is limited
by the encounter of S with E. This ratio approaches 10 8 to 10 9 M -1 s _1 , the fastest rate at
which two uncharged solutes can approach each other by diffusion at physiological
temperature. Enzymes that can catalyze reactions at this extremely rapid rate are dis-
cussed in Section 6.4.

The k cat /K m ratio is useful for comparing the activities of different enzymes. It is
also possible to assess the efficiency of an enzyme by measuring its catalytic proficiency.
This value is equal to the rate constants for a reaction in the presence of the enzyme
( k cat /K m ) divided by the rate constant for the same reaction in the absence of the en-
zyme (fc n ). Surprisingly few catalytic proficiency values are known because most chemi-
cal reactions occur extremely slowly in the absence of enzymes — so slowly that their
nonenzymatic rates are very difficult to measure. The reaction rates are often measured
in special steel-enclosed glass vessels at temperatures in excess of 300°C.

Table 5.2 lists several examples of known catalytic proficiencies. Typical values
range from 10 14 to 10 20 but some are quite a bit higher (up to 10 24 ). The current record
holder is uroporphyrinogen decarboxylase, an enzyme required for a step in the por-
phyrin synthesis pathway. The difficulty in obtaining rate constants for nonenzymatic
reactions is illustrated by the half-life for the uncatalyzed reaction — about 2 billion
years! The catalytic proficiency values in Table 5.2 emphasize one of the main properties
of enzymes, namely, their ability to increase the rates of reactions that would normally
occur too slowly to be useful.

5.5 Measurement of K m and V max

The kinetic parameters of an enzymatic reaction can provide valuable information about
the specificity and mechanism of the reaction. The key parameters are K m and V max
because fc cat can be calculated if V max is known.

146 CHAPTER 5 Properties of Enzymes

Table 5.2 Catalytic proficiencies of some enzymes

Nonenzymatic
rate constant
(fc„ in s' 1 )

Enzymatic rate
constant ( k cat /K m
in M 's 1 )

Catalytic

proficiency

Carbonic anhydrase

10" 1

7 X 10 6

7 X 10 7

Chymotrypsin

4 x icr 9

9 X 10 7

2 X 10 16

Chorismate mutase

1(T 5

2 X 10 6

2 X 10 11

Triose phosphate isomerase

4 x icr 6

4 X 10 8

10 14

Cytidine deaminase

10 -i°

3 X 10 6

3 X 10 16

Adenosine deaminase

2 X 1(T 10

10 7

5 X 10 16

Mandelate racemase

3 x icr 13

10 6

3 X 10 18

/3-Amylase

7 X 1(T 14

10 7

10 20

Fumarase

icr 13

1 0 9

10 21

Arginine decarboxylase

9 x icr 16

10 6

10 21

Alkaline phosphatase

icr 15

3 X 10 7

3 X 10 22

Orotidine 5'-phosphate
decarboxylase

3 x icr 16

6 x 10 7

2 X 10 23

Uroporphyrinogen

decarboxylase

1 o -17

2 X 10 7

2 X 10 24

K m and V max for an enzyme -catalyzed reaction can be determined in several ways.
Both values can be obtained by the analysis of initial velocities at a series of substrate
concentrations and a fixed concentration of enzyme. In order to obtain reliable values
for the kinetic constants the [S] points must be spread out both below and above K m to
produce a hyperbola. It is difficult to determine either K m or V max directly from a graph

▲ Maximum catalytic proficiency. Uropor-
phyrinogen decarboxylase is the current
record holder for maximum catalytic profi-
ciency. It catalyzes a step in the heme syn-
thesis pathway. The enzyme shown here is a
human (Homo sapiens) variant with a bound
porphoryrin molecule at the active site of
each monomer. [PDB 2Q71]

BOX 5.2 HYPERBOLAS VERSUS STRAIGHT LINES

We have seen that a plot of substrate concentration ([S])
versus the initial velocity of a reaction (v 0 ) produces a hy-
perbolic curve as shown in Figures 5.4 and 5.5. The general
equation for a rectangular hyperbola (Equation 5.12) and
the Michaelis-Menten equation have the same form
(Equation 5.13).

Its very difficult to determine V max from a plot of enzyme
kinetic data since the hyperbolic curve that shows the relation-
ship between substrate concentration and initial velocity is as-
ymptotic to V max and it is experimentally difficult to achieve
the concentration of substrate required to estimate V max . For
these reasons, it is often easier to convert the hyperbolic curve
to a linear form that matches the general formula y - mx + b,
where m is the slope of the line and b is the y-axis intercept.
The first step in transforming the original Michaelis-Menten
equation to this general form of a linear equation is to invert
the terms so that the K m + [S] term is on top of the right-hand
side. This is done by taking the reciprocal of each side — a
transformation that will be familiar to many who are familiar
with hyperbolic curves.

The next two steps involve separating terms and cancel-
ing [S] in the second term on the right-hand side of the
equation. This form of the Michaelis-Menten equation is
called the Lineweaver-Burk equation and it resembles the
general form of a linear equation, y - mx + b , where y is the
reciprocal of v 0 and x values are the reciprocal of [S]. A plot
of data in this form is referred to as a double-reciprocal plot.
The slope of the line will be K m /V max and the y-axis intercept
will be W max .

The original reason for this sort of transformation was
to calculate K m and V max from experimental data. It was eas-
ier to plot the reciprocal values of v 0 and [S] and draw a
straight line through the points in order to calculate the ki-
netic constants. Nowadays, there are computer programs that
can accurately fit the data to a hyperbolic curve and calculate
the constants so the Lineweaver-Burk plot is no longer nec-
essary for this type of analysis. In this book we will still use
the Lineweaver-Burk plots to illustrate some general features
of enzyme kinetics but they are rarely used for their original
purpose of data analysis.

1 = K m + [S] ^ = K m + [S] = ^ I< m b 1 +

v o y max [s] v 0 y max [s] v max [S] v 0 kmax [S] V m

5.6 Kinetics of Multisubstrate Reactions 147

of initial velocity versus concentration because the curve approaches V max asymptoti-
cally. However, accurate values can be determined by using a suitable computer pro-
gram to fit the experimental results to the equation for the hyperbola.

The Michaelis-Menten equation can be rewritten in order to obtain values for V max
and K m from straight lines on graphs. The most commonly used transformation is the
double-reciprocal, or Lineweaver-Burk, plot in which the values of l/v 0 are plotted
against 1/[S] (Figure 5.6 ). The absolute value of 1 /K m is obtained from the intercept of
the line at the x axis, and the value of 1/V max is obtained from the y intercept. Although
double-reciprocal plots are not the most accurate methods for determining kinetic con-
stants, they are easily understood and provide recognizable patterns for the study of en-
zyme inhibition, an extremely important aspect of enzymology that we will examine
shortly.

Values of fc cat can be obtained from measurements of V max only when the absolute
concentration of the enzyme is known. Values of K m can be determined even when en-
zymes have not been purified provided that only one enzyme in the impure preparation
can catalyze the observed reaction.

Lineweaver-Burk equation:

J_ = (^m|l + _J_

5.6 Kinetics of Multisubstrate Reactions

Until now, we have only been considering reactions where a single substrate is con-
verted to a single product. Let’s consider a reaction in which two substrates, A and B, are
converted to products P and Q.

▲ Figure 5.6

Double-reciprocal (Lineweaver-Burk) plot.

This plot is derived from a linear transforma-
tion of the Michaelis-Menten equation.
Values of 1/vq are plotted as a function of
1/[S] values.

E + A + B (EAB) -> E + P + Q (5.27)

Kinetic measurements for such multisubstrate reactions are a little more complicated
than simple one-substrate enzyme kinetics. For many purposes, such as designing an
enzyme assay, it’s sufficient simply to determine the K m for each substrate in the pres-
ence of saturating amounts of each of the other substrates as we described for chemi-
cal reactions (Section 5.2A). The simple enzyme kinetics discussed in this chapter can
be extended to distinguish among several mechanistic possibilities for multisubstrate
reactions, such as group transfer reactions. This is done by measuring the effect of
variations in the concentration of one substrate on the kinetic results obtained for the
other.

Multisubstrate reactions can occur by several different kinetic schemes. These
schemes are called kinetic mechanisms because they are derived entirely from kinetic
experiments. Kinetic mechanisms are commonly represented using the notation intro-
duced by W. W. Cleland. The sequence of steps proceeds from left to right (Figure 5.7).
The addition of substrate molecules (A, B, C, . . .) to the enzyme and the release of
products (P, Q, R, . . .) from the enzyme are indicated by arrows pointing toward
(substrate binding) or from (product release) the line. The various forms of the en-
zyme (free E, ES complexes, or EP complexes) are written under a horizontal line. The
ES complexes that undergo chemical transformation when the active site is filled are
shown in parentheses.

Sequential reactions (Figure 5.7a) require all the substrates to be present before any
product is released. Sequential reactions can be either ordered, with an obligatory order
for the addition of substrates and release of products, or random. In ping-pong reactions
(Figure 5.7b), a product is released before all the substrates are bound. In a bisubstrate
ping-pong reaction, the first substrate is bound, the enzyme is altered by substitution,
and the first product is released. Then the second substrate is bound, the altered enzyme
is restored to its original form, and the second product is released. A ping-pong mecha-
nism is sometimes called a substituted-enzyme mechanism because of the covalent
binding of a portion of a substrate to the enzyme. The binding and release of ligands in
a ping-pong mechanism are usually indicated by slanted lines. The two forms of the en-
zyme are represented by E (unsubstituted) and F (substituted).

148

CHAPTER 5 Properties of Enzymes

Irreversible inhibitors are described in
Section 5.8.

KEY CONCEPT

Reversible inhibitors bind to enzymes
and either prevent substrate binding or
block the reaction leading to formation
of product.

(a) Sequential reactions

A B P Q

A A

V T

E EA (EAB) (EPQ) EQ E

Ordered

A B P Q

i eb i i ep i

BA Q P

Random

(b) Ping-pong reaction

E (EA)(FP) F (FB)(EQ) E

▲ Figure 5.7

Notation for bisubstrate reactions, (a) In sequential reactions, all substrates are bound before a product
is released. The binding of substrates may be either ordered or random, (b) In ping-pong reactions,
one substrate is bound and a product is released, leaving a substituted enzyme. A second substrate
is then bound and a second product released, restoring the enzyme to its original form.

5.7 Reversible Enzyme Inhibition

An enzyme inhibitor (I) is a compound that binds to an enzyme and interferes with its
activity. Inhibitors can act by preventing the formation of the ES complex or by block-
ing the chemical reaction that leads to the formation of product. As a general rule,
inhibitors are small molecules that bind reversibly to the enzyme they inhibit. Cells
contain many natural enzyme inhibitors that play important roles in regulating me-
tabolism. Artificial inhibitors are used experimentally to investigate enzyme mecha-
nisms and decipher metabolic pathways. Some drugs, and many poisons, are enzyme
inhibitors.

Some inhibitors bind covalently to enzymes causing irreversible inhibition but
most biologically relevant inhibition is reversible. Reversible inhibitors are bound to
enzymes by the same weak, noncovalent forces that bind substrates and products.
The equilibrium between free enzyme (E) plus inhibitor (I) and the El complex is
characterized by a dissociation constant. In this case, the constant is called the
inhibition constant,^.

E + | — El K d = K; = ^ (5.28)

The basic types of reversible inhibition are competitive, uncompetitive, noncom-
petitive and mixed. These can be distinguished experimentally by their effects on the ki-
netic behavior of enzymes (Table 5.3). Figure 5.8 shows diagrams representing modes
of reversible enzyme inhibition.

5.7 Reversible Enzyme Inhibition 149

Table 5.3 Effects of reversible inhibitors on kinetic constants

Type of inhibitor

Effect

Competitive (1 binds to E only)

Raises K m

V max remains unchanged

Uncompetitive (1 binds to ES only)

Lowers V max and K m

Ratio of V max /K m remains unchanged

Noncompetitive (1 binds to E or ES)

Lowers V max
K m remains unchanged

A. Competitive Inhibition

Competitive inhibitors are the most commonly encountered inhibitors in biochem-
istry. In competitive inhibition, the inhibitor can bind only to free enzyme molecules
that have not bound any substrate. Competitive inhibition is illustrated in Figure 5.8
and by the kinetic scheme in Figure 5.9a. In this scheme only ES can lead to the for-
mation of product. The formation of an El complex removes enzyme from the nor-
mal pathway.

Once a competitive inhibitor is bound to an enzyme molecule, a substrate mole-
cule cannot bind to that enzyme molecule. Conversely, the binding of substrate to an
enzyme molecule prevents the binding of an inhibitor. In other words, S and I compete
for binding to the enzyme molecule. Most commonly, S and I bind at the same site on
the enzyme, the active site. This type of inhibition is termed classical competitive inhi-
bition (Figure 5.8). This is not the only kind of competitive inhibition (see Figure 5.8).
In some cases, such as allosteric enzymes (Section 5.10), the inhibitor binds at a differ-
ent site and this alters the substrate binding site preventing substrate binding. This
type of inhibition is called nonclassical competitive inhibition. When both I and S are

(a) Classical competitive inhibition (b) Nonclassical competitive inhibition

co = db - p- 1 *!

▲ Competitive inhibition. The active
ingredient in the weed killer Roundup® is
glyphosate, a competitive inhibitor of the
plant enzyme 5-enolpyruvylshikimate-3-
phosphate synthase. (See Box 17.2 in
Chapter 17.)

The substrate (S) and the inhibitor The binding of substrate (S) at the active

(I) compete for the same site on site prevents the binding of inhibitor (I)

the enzyme. at a separate site and vice versa.

(d) Noncompetitive inhibition

The inhibitor (I) binds only to the
enzyme substrate (ES) complex
preventing the conversion of
substrate (S) to product.

The inhibitor (I) can bind to either E or
ES. The enzyme becomes inactive when
I binds. Substrate (S) can still bind to
the El complex but conversion to
product is inhibited.

◄ Figure 5.8

Diagrams of reversible enzyme inhibition. In

this scheme, catalytically competent enzymes
are green and inactive enzymes are red.

150

CHAPTER 5 Properties of Enzymes

(a)

k i

E + S < » ES

+ ^-i

E + P

▲ Figure 5.9

Competitive inhibition, (a) Kinetic scheme illustrating the binding of I to E. Note that this is an ex-
pansion of Equation 5.11 that includes formation of the El complex, (b) Double-reciprocal plot. In
competitive inhibition, l/ max remains unchanged and K m increases. The black line labeled “Control”
is the result in the absence of inhibitor. The red lines are the results in the presence of inhibitor,
with the arrow showing the direction of increasing [I].

▲ Ibuprofen, the active ingredient in many
over-the-counter painkillers, is a competitive
inhibitor of the enzyme cyclooxygenase
(COX). (See Box 16.1 Chapter 16.)

coo e

C H2

CH 2

COO 0

Succinate

COO'

< f H2 ,

coo'

,©

Malonate

present in a solution, the proportion of the enzyme that is able to form ES complexes
depends on the concentrations of substrate and inhibitor and their relative affinities
for the enzyme.

The amount of El can be reduced by increasing the concentration of S. At suffi-
ciently high concentrations the enzyme can still be saturated with substrate. Therefore,
the maximum velocity is the same in the presence or in the absence of an inhibitor.
The more competitive inhibitor present, the more substrate needed for half- saturation.
We have shown that the concentration of substrate at half- saturation is K m . In the pres-
ence of increasing concentrations of a competitive inhibitor, K m increases. The new
value is usually referred to as the apparent (X^ p ). On a double-reciprocal plot,
adding a competitive inhibitor shows as a decrease in the absolute value of the intercept
at the x axis 1 /K m , whereas the y intercept VV max remains the same (Figure 5.9b).

Many classical competitive inhibitors are substrate analogs — compounds that are
structurally similar to substrates. The analogs bind to the enzyme but do not react.
For example, the enzyme succinate dehydrogenase converts succinate to fumarate
(Section 13.3#6). Malonate resembles succinate and acts as a competitive inhibitor of
the enzyme.

B. Uncompetitive Inhibition

Uncompetitive inhibitors bind only to ES and not to free enzyme (Figure 5.10a). In
uncompetitive inhibition, V max is decreased (W max is increased) by the conversion of some
molecules of E to the inactive form ESI. Since it is the ES complex that binds I, the de-
crease in V max is not reversed by the addition of more substrate. Uncompetitive in-
hibitors also decrease the K m (seen as an increase in the absolute value of 1 /K m on a
double- reciprocal plot) because the equilibria for the formation of both ES and ESI are
shifted toward the complexes by the binding of I. Experimentally, the lines on a double-
reciprocal plot representing varying concentrations of an uncompetitive inhibitor all
have the same slope indicating proportionally decreased values for K m and U max (Figure
5.10b). This type of inhibition usually occurs only with multisubstrate reactions.

C. Noncompetitive Inhibition

Noncompetitive inhibitors can bind to E or ES forming inactive El or ESI complexes, re-
spectively (Figure 5.11a). These inhibitors are not substrate analogs and do not bind at
the same site as S. The classic case of noncompetitive inhibition is characterized by an

5.7 Reversible Enzyme Inhibition 151

(a)

E + S < > ES > E + P

K i

ESI

(a)

E + S ES

+ +

I I

El + S ESI

E + P

apparent decrease in V max ( W max appears to increase) with no change in K m . On a
double-reciprocal plot, the lines for classic noncompetitive inhibition intersect at the
point on the x axis corresponding to 1 /K m (Figure 5.1 lb). The common x-axis intercept
indicates that K m isn’t affected. The effect of noncompetitive inhibition is to reversibly
titrate E and ES with I removing active enzyme molecules from solution. This inhibi-
tion cannot be overcome by the addition of S. Classic noncompetitive inhibition is rare
but examples are known among allosteric enzymes. In these cases, the noncompetitive
inhibitor probably alters the conformation of the enzyme to a shape that can still bind S
but cannot catalyze any reaction.

Most enzymes do not conform to the classic form of noncompetitive inhibition
where K m is unchanged. In most cases, both K m and V max are affected because the affin-
ity of the inhibitor for E is different than its affinity for ES. These cases are often referred
to as mixed inhibition (Figure 5.12).

D. Uses of Enzyme Inhibition

Reversible enzyme inhibition provides a powerful tool for probing enzyme activity. In-
formation about the shape and chemical reactivity of the active site of an enzyme can be
obtained from experiments involving a series of competitive inhibitors with systemati-
cally altered structures.

The pharmaceutical industry uses enzyme inhibition studies to design clinically
useful drugs. In many cases, a naturally occurring enzyme inhibitor is used as the start-
ing point for drug design. Instead of using random synthesis and testing of potential in-
hibitors, some investigators are turning to a more efficient approach known as rational
drug design. Theoretically, with the greatly expanded bank of knowledge about enzyme
structure, inhibitors can now be rationally designed to fit the active site of a target
enzyme. The effects of a synthetic compound are tested first on isolated enzymes and
then in biological systems. However, even if a compound has suitable inhibitory activ-
ity, other problems may be encountered. For example, the drug may not enter the target
cells, may be rapidly metabolized to an inactive compound, may be toxic to the host or-
ganism, or the target cell may develop resistance to the drug.

◄ Figure 5.10

Uncompetitive inhibition, (a) Kinetic scheme
illustrating the binding of I to ES.

(b) Double-reciprocal plot. In uncompetitive
inhibition, both l/ max and K m decrease (i.e.,
the absolute values of both l/l/ max and 1/K m
obtained from they and x intercepts,
respectively, increase). The ratio KJ l/ max ,
the slope of the lines, remains unchanged.

◄ Figure 5.1 1

Classic noncompetitive inhibition, (a) Kinetic
scheme illustrating the binding of I to E
or ES. (b) Double-reciprocal plot. F max
decreases, but K m remains the same.

[s]

▲ Figure 5.12

Double-reciprocal plot showing mixed Inhibi-
tion. Both y max and K m are affected when
the inhibitor binds with different affinities to
E and ES.

152 CHAPTER 5 Properties of Enzymes

(b)

h 2 n

▲ Figure 5.13

Comparison of a substrate and a designed in-
hibitor of purine nucleoside phosphorylase.

The two substrates of this enzyme are
guanosine and inorganic phosphate, (a)
Guanosine. (b) A potent inhibitor of the en-
zyme. N-9 of guanosine has been replaced
by a carbon atom. The chlorinated benzene
ring binds to the sugar-binding site of the
enzyme, and the acetate side chain binds to
the phosphate-binding site.

The advances made in drug synthesis are exemplified by the design of a series of in-
hibitors of the enzyme purine nucleoside phosphorylase. This enzyme catalyzes a
degradative reaction between phosphate and the nucleoside guanosine whose structure
is shown in Figure 5.13a. With computer modeling, the structures of potential in-
hibitors were designed and fit into the active site of the enzyme. One such compound
(Figure 5.13b) was synthesized and found to be 100 times more inhibitory than any
compound made by the traditional trial- and-error approach. Researchers hope that the
rational design approach will produce a drug suitable for treating autoimmune disor-
ders such as rheumatoid arthritis and multiple sclerosis.

5.8 Irreversible Enzyme Inhibition

In contrast to a reversible enzyme inhibitor, an irreversible enzyme inhibitor forms a
stable covalent bond with an enzyme molecule thus removing active molecules from
the enzyme population. Irreversible inhibition typically occurs by alkylation or acylation
of the side chain of an active-site amino acid residue. There are many naturally occur-
ring irreversible inhibitors as well as the synthetic examples described here.

An important use of irreversible inhibitors is the identification of amino acid
residues at the active site by specific substitution of their reactive side chains. In this
process, an irreversible inhibitor that reacts with only one type of amino acid is in-
cubated with a solution of enzyme that is then tested for loss of activity. Ionizable
side chains are modified by acylation or alkylation reactions. For example, free
amino groups such as the e-amino group of lysine react with an aldehyde to form a
Schiff base that can be stabilized by reduction with sodium borohydride (NaBH 4 )
(Figure 5.14).

The nerve gas diisopropyl fluorophosphate (DFP) is one of a group of organic
phosphorus compounds that inactivate hydrolases with a reactive serine as part of the
active site. These enzymes are called serine proteases or serine esterases, depending on
their reaction specificity. The serine protease chymotrypsin, an important digestive
enzyme, is inhibited irreversibly by DFP (Figure 5.15). DFP reacts with the serine
residue at chymotrypsin’s active site (Ser-195) to produce diisopropylphosphoryl-
chymotrypsin.

Some organophosphorus inhibitors are used in agriculture as insecticides; others,
such as DFP, are useful reagents for enzyme research. The original organophosphorus
nerve gases are extremely toxic poisons developed for military use. The major biological
action of these poisons is irreversible inhibition of the serine esterase acetyl-
cholinesterase that catalyzes hydrolysis of the neurotransmitter acetylcholine. When
acetylcholine released from an activated nerve cell binds to its receptor on a second
nerve cell, it triggers a nerve impulse. The action of acetylcholinesterase restores the cell
to its resting state. Inhibition of this enzyme can cause paralysis.

Lys

(CH 2 ) 4

h 2 o

Lys

(CH 2 ) 4

Lys

(CH 2 ) 4

nh 2

NaBH 4

h 2 o

Schiff base

▲ Figure 5.14

Reaction of the e-amino group of a lysine residue with an aldehyde. Reduction of the Schiff base with
sodium borohydride (NaBH 4 ) forms a stable substituted enzyme.

5.9 Regulation of Enzyme Activity 153

Figure 5.15 ►

Irreversible Inhibition by DFP. Diisopropyl fluorophosphate (DFP) reacts with a single, highly nucle-
ophilic serine residue (Ser-195) at the active site of chymotrypsin, producing inactive diisopropyl-
phosphoryl-chymotrypsin. DFP inactivates serine proteases and serine esterases.

Ser-195

5.9 Regulation of Enzyme Activity

At the beginning of this chapter, we listed several advantages to using enzymes as catalysts
in biochemical reactions. Clearly, the most important advantage is to speed up reactions
that would otherwise take place too slowly to sustain life. One of the other advantages of
enzymes is that their catalytic activity can be regulated in various ways. The amount of
an enzyme can be controlled by regulating the rate of its synthesis or degradation. This
mode of control occurs in all species but it often takes many minutes or hours to
synthesize new enzymes or to degrade existing enzymes.

In all organisms, rapid control — on the scale of seconds or less — can be accom-
plished through reversible modulation of the activity of regulated enzymes. In this con-
text, we define regulated enzymes as those enzymes whose activity can be modified in a
manner that affects the rate of an enzyme -catalyzed reaction. In many cases, these regu-
lated enzymes control a key step in a metabolic pathway. The activity of a regulated en-
zyme changes in response to environmental signals, allowing the cell to respond to
changing conditions by adjusting the rates of its metabolic processes.

In general, regulated enzymes become more active catalysts when the concentra-
tions of their substrates increase or when the concentrations of the products of their
metabolic pathways decrease. They become less active when the concentrations of their
substrates decrease or when the products of their metabolic pathways accumulate. Inhi-
bition of the first enzyme unique to a pathway conserves both material and energy by
preventing the accumulation of intermediates and the ultimate end product. The activity
of regulated enzymes can be controlled by noncovalent allosteric modulation or covalent
modification.

Allosteric enzymes are enzymes whose properties are affected by changes in struc-
ture. The structural changes are mediated by interaction with small molecules. We saw
an example of allostery in the previous chapter when we examined the binding of oxygen
to hemoglobin. Allosteric enzymes often do not exhibit typical Michaelis-Menten kinet-
ics due to cooperative binding of substrate, as is the case with hemoglobin.

Figure 5.16 shows a v 0 versus [S] curve for an allosteric enzyme with cooperative
binding of substrate. Sigmoidal curves result from the transition between two states of
the enzyme. In the absence of substrate, the enzyme is in the T state. The conformation
of each subunit is in a shape that binds substrate inefficiently and the rate of the reac-
tion is slow. As substrate concentration is increased, enzyme molecules begin to bind
substrate even though the affinity of the enzyme in the T state is low. When a subunit
binds substrate, the enzyme undergoes a conformational change that converts the en-
zyme to the R state and the reaction takes place. The kinetic properties of the enzyme
subunit in the T state and the R state are quite different — each conformation by itself
could exhibit standard Michaelis-Menten kinetics.

The conformational change in the subunit that initially binds a substrate molecule
affects the other subunits in the multisubunit enzyme. The conformations of these
other subunits are shifted toward the R state where their affinity for substrate is much
higher. They can now bind substrate at a much lower concentration than when they
were in the T state.

Allosteric phenomena are responsible for the reversible control of many regulated
enzymes. In Section 4.13C, we saw how the conformation of hemoglobin and its affinity
for oxygen change when 2,3-frisphosphoglycerate is bound. Many regulated enzymes
also undergo allosteric transitions between active (R) states and inactive (T) states.
These enzymes have a second ligand-binding site away from their catalytic centers
called the regulatory site or allosteric site. An allosteric inhibitor or activator, also called an
allosteric modulator or allosteric effector, binds to the regulatory site and causes a con-
formational change in the regulated enzyme. This conformational change is transmitted

h 3 C

H — C — O

H 3 C

-P

11^

C H 3

o — c — H

ch 3

Diisopropyl fluorophosphate
(DFP)

^H®

Ser-195

CH 2

h 3 c O ch 3

I \ /-> I

H— C— O— P— O— C— H

H,C

CH 3

Ser-195

H 3 C O ch 3

I I I

H— C— O— P— O— C— H

H 3 C O ch 3
Diisopropylphosphoryl-chymotrypsin

Aspartate transcarbamoylase (ATCase),
another well-characterized allosteric
enzyme, is described in Chapter 18.

[S]

▲ Figure 5.16

Cooperativity. Plot of initial velocity as a
function of substrate concentration for an
allosteric enzyme exhibiting cooperative
binding of substrate.

154 CHAPTER 5 Properties of Enzymes

KEY CONCEPT

Allosteric enzymes often have multiple
subunits and substrate binding is
cooperative. This produces a sigmoidal
curve when velocity is plotted against
substrate concentration.

ch 2 oh

c=o

HO — C — H

H — C — OH

I ©

ch 2 opo 3 ^

Fructose 6-phosphate

ATP ADP

Phosphofructokinase - 1

ch 2 opo 3 ®

c=o

HO — C — H

+ H®

H — C — OH + M

H — C — OH

I ©

ch 2 opo 3 ^

Fructose 1,6-b/sphosphate

▲ Figure 5.17

Reaction catalyzed by phosphofructokinase-1.

to the active site of the enzyme, which changes shape sufficiently to alter its activity. The
regulatory and catalytic sites are physically distinct regions of the protein — usually lo-
cated on separate domains and sometimes on separate subunits. Allosterically regulated
enzymes are often larger than other enzymes.

First, we examine an enzyme that undergoes allosteric (noncovalent) regulation
and then we list some general properties of such enzymes. Next, we describe two models
that explain allosteric regulation in terms of changes in the conformation of regulated
enzymes. Finally, we discuss a closely related group of regulatory enzymes — those subject
to covalent modification.

coo°

C — OPO,®

ch 2

▲ Figure 5.18

Phosphoenolpyruvate. This intermediate of
glycolysis is an allosteric inhibitor of phos-
phofructokinase- 1 from Escherichia coli.

A. Phosphofructokinase Is an Allosteric Enzyme

Bacterial phosphofructokinase-1 ( Escherichia coli) provides a good example of allosteric
inhibition and activation. Phosphofructokinase-1 catalyzes the ATP-dependent phos-
phorylation of fructose 6-phosphate to produce fructose 1,6-frzsphosphate and ADP
(Figure 5.17). This reaction is one of the first steps of glycolysis, an ATP-generating
pathway for glucose degradation described in detail in Chapter 11. Phosphoenolpyruvate
(Figure 5.18), an intermediate near the end of the glycolytic pathway, is an allosteric
inhibitor of E. coli phosphofructokinase-1. When the concentration of phospho-
enolpyruvate rises, it indicates that the pathway is blocked beyond that point. Further
production of phosphoenolpyruvate is prevented by inhibiting phosphofructokinase- 1
(see feedback inhibition, Section 10.2C).

ADP is an allosteric activator of phosphofructokinase-1. This may seem strange from
looking at Figure 5.17 but keep in mind that the overall pathway of glycolysis results in net
synthesis of ATP from ADR Rising ADP levels indicate a deficiency of ATP and glycolysis
needs to be stimulated. Thus, ADP activates phosphofructokinase-1 in spite of the fact
that ADP is a product in this particular reaction.

Phosphoenolpyruvate and ADP affect the binding of the substrate fructose 6-phos-
phate to phosphofructokinase-1. Kinetic experiments have shown that there are four
binding sites on phosphofructokinase-1 for fructose 6-phosphate and structural experi-
ments have confirmed that E. coli phosphofructokinase-1 (M r 140,000) is a tetramer
consisting of four identical subunits. Figure 5.19 shows the structure of the enzyme
complexed with its products, fructose 1,6-fcphosphate and ADP, and a second mole-
cule of ADP, an allosteric activator. Two of the subunits shown in Figure 5.19a associate
to form a dimer. The two products are bound in the active site located between two do-
mains of each chain — ADP is bound to the large domain and fructose 1,6-frisphosphate
is bound mostly to the small domain. Two of these dimers interact to form the complete
tetrameric enzyme.

A notable feature of the structure of phosphofructokinase-1 (and a general feature
of regulated enzymes) is the physical separation of the active site and the regulatory

5.9 Regulation of Enzyme Activity 155

site on each subunit. (In some regulated enzymes the active sites and regulatory sites
are on different subunits.) The activator ADP binds at a distance from the active site in
a deep hole between the subunits. When ADP is bound to the regulatory site, phospho-
fructokinase-1 assumes the R conformation, which has a high affinity for fructose 6-
phosphate. When the smaller compound phosphoenolpyruvate is bound to the same
regulatory site the enzyme assumes a different conformation, the T conformation,
which has a lower affinity for fructose 6-phosphate. The transition between conforma-
tions is accomplished by a slight rotation of one rigid dimer relative to the other. The
cooperativity of substrate binding is tied to the concerted movement of an arginine
residue in each of the four fructose 6-phosphate binding sites located near the inter-
face between the dimers. Movement of the side chain of this arginine from the active
site lowers the affinity for fructose 6-phosphate. In many organisms, phosphofructoki-
nase-1 is larger and is subject to more complex allosteric regulation than in E. coli as
you will see in Chapter 1 1 .

Activators can affect either V max or K m or both. Its important to recognize that the
binding of an activator alters the structure of an enzyme and this alteration converts it
to a different form that may have quite different kinetic properties. In most cases, the
differences between the kinetic properties of the R and T forms are more complex than
the differences we saw with enzyme inhibitors in Section 5.7.

B. General Properties of Allosteric Enzymes

Examination of the kinetic and physical properties of allosteric enzymes has shown that
they have the following general features:

1. The activities of allosteric enzymes are changed by metabolic inhibitors and activa-
tors. Often these allosteric effectors do not resemble the substrates or products of
the enzyme. For example, phosphoenolpyruvate (Figure 5.18) resembles neither
the substrate nor the product (Figure 5.17) of phosphofructokinase. Consideration
of the structural differences between substrates and metabolic inhibitors originally
led to the conclusion that allosteric effectors are bound to regulatory sites separate
from catalytic sites.

2. Allosteric effectors bind noncovalently to the enzymes they regulate. (There is a
special group of regulated enzymes whose activities are controlled by covalent
modification, described in Section 5.10D.) Many effectors alter the K m of the en-
zyme for a substrate; but some alter the V max . Allosteric effectors themselves are not
altered chemically by the enzyme.

3. With few exceptions, regulated enzymes are multisubunit proteins. (But not all
multisubunit enzymes are regulated.) The individual polypeptide chains of a
regulated enzyme may be identical or different. For those with identical sub-
units (such as phosphofructokinase- 1 from E. coli), each polypeptide chain can
contain both the catalytic and regulatory sites and the oligomer is a symmetric
complex, most often possessing two or four protein chains. Regulated enzymes
composed of nonidentical subunits have more complex, but usually symmetric,
arrangements.

4. An allosterically regulated enzyme usually has at least one substrate for which the
v 0 versus [S] curve is sigmoidal rather than hyperbolic (Section 5.9). Phospho-
fructokinase- 1 exhibits Michaelis-Menten (hyperbolic) kinetics with respect to
one substrate, ATP, but sigmoidal kinetics with respect to its other substrate, fruc-
tose 6-phosphate. A sigmoidal curve is caused by positive cooperativity of sub-
strate binding and this is made possible by the presence of multiple substrate
binding sites in the enzyme — four binding sites in the case of tetrameric phospho-
fructokinase- 1.

The allosteric R v T transition between the active and the inactive conformations
of a regulatory enzyme is rapid. The ratio of R to T is controlled by the concentrations of
the various ligands and the relative affinities of each conformation for these ligands. In
the simplest cases, substrate and activator molecules bind only to enzyme in the R state
(Er) and inhibitor molecules bind only to enzyme in the T state (E T ).

KEY CONCEPT

Allosteric effectors shift the concentra-
tions of the R and T forms of an allosteric
enzyme.

(a)

▲ Figure 5.19

The R conformation of phosphofructokinase-1
from E. coli. The enzyme is a tetramer of
identical chains, (a) Single subunit, shown
as a ribbon. The products, fructose 1,6-
b/sphosphate (yellow) and ADP (green), are
bound in the active site. The allosteric acti-
vator ADP (red) is bound in the regulatory
site, (b) Tetramer. Two are blue, and two are
purple. The products, fructose 1,6-
b/'sphosphate (yellow) and ADP (green), are
bound in the four active sites. The allosteric
activator ADP (red) is bound in the four reg-
ulatory sites, at the interface of the sub-
units. [PDB 1PFK].

The relationship between the regula-
tion of an individual enzyme and a
pathway is discussed in Section 10.2B,
where we encounter terms such as
feedback inhibition and feedforward
activation.

156 CHAPTER 5 Properties of Enzymes

Figure 5.20 ►

Role of cooperativity of binding in regulation.

The activity of an allosteric enzyme with a
sigmoidal binding curve can be altered
markedly when either an activator or an in-
hibitor is bound to the enzyme. Addition of
an activator can lower the apparent K m rais-
ing the activity at a given [S]. Conversely,
addition of an inhibitor can raise the appar-
ent K m producing less activity at a given [S].

Allosteric

transition

; >

(5.29)

E t

(5.30)

These simplified examples illustrate the main property of allosteric effectors — they shift
the steady- state concentrations of free Ej and E R .

Figure 5.20 illustrates the regulatory role that cooperative binding can play. Addi-
tion of an activator can shift the sigmoidal curve toward a hyperbolic shape, lowering
the apparent K m (the concentration of substrate required for half- saturation) and rais-
ing the activity at a given [S]. The addition of an inhibitor can raise the apparent K m of
the enzyme and lower its activity at any particular concentration of substrate.

The addition of S leads to an increase in the concentration of enzyme in the R con-
formation. Conversely, the addition of inhibitor increases the proportion of the T
species. Activator molecules bind preferentially to the R conformation leading to an
increase in the R/T ratio. Note that this simplified scheme does not show that there are
multiple interacting binding sites for both S and I.

Some allosteric inhibitors are nonclassical competitive inhibitors (Figure 5.8). For
example, Figure 5.20 describes an enzyme that has a higher apparent K m for its sub-
strate in the presence of the allosteric inhibitor but an unaltered V max . Therefore, the
allosteric modulator is a competitive inhibitor.

Some regulatory enzymes exhibit noncompetitive inhibition patterns where bind-
ing of a modulator at the regulatory site does not prevent substrate from binding but
appears to distort the conformation of the active site sufficiently to decrease the activity
of the enzyme.

C. Two Theories of Allosteric Regulation

Recall that most proteins are made up of two or more polypeptide chains (Section 4.8).
Enzymes are typical proteins — most of them have multiple subunits. This complicates
our understanding of regulation. There are two general models that explain the cooper-
ative binding of ligands to multimeric proteins. Both models describe the cooperative
transitions in simple quantitative terms.

The concerted model, or symmetry model, was devised to explain the cooperative
binding of identical ligands, such as substrates. It was first proposed in 1965 by

5.9 Regulation of Enzyme Activity

157

▲ Figure 5.21

Two models for cooperativity of binding of substrate (S) to a tetrameric protein. A two-subunit protein is shown for simplicity. In all cases, the enzymati-
cally active subunit (R) is colored green and the inactive conformation (T) is colored red. (a) In the simplified concerted model, both subunits are ei-
ther in the R conformation or the T conformation. Substrate (S) can bind to subunits in either conformation but binding to T is assumed to be weaker
than binding to R. Cooperativity is explained by postulating that when substrate binds to a subunit in the T conformation (red), it shifts the protein
into a conformation where both subunits are in the R conformation, (b) In the sequential model, one subunit may be in the R conformation while an-
other is in the T conformation. As in the concerted model, both conformations can bind substrate. Cooperativity is achieved by postulating that sub-
strate binding causes the subunit to shift to the R conformation and that when one subunit has adopted the R conformation, the other one is more
likely to bind substrate and undergo a conformation change (diagonal lines).

Jacques Monod, Jeffries Wyman, and Jean-Pierre Changeux and it’s sometimes known
as the MWC model. The concerted model assumes there is one substrate binding site
on each subunit. According to the concerted model, the conformation of each subunit
is constrained by its association with other subunits and when the protein changes
conformation it retains its molecular symmetry (Figure 5.21a). Thus, there are two
conformations in equilibrium, R and T. When a subunit is in the R conformation it
has a high affinity for the substrate. Subunits in the T conformation have a low affin-
ity for the substrate. The binding of substrate to one subunit shifts the equilibrium
since it “locks” the other subunits in the R conformation making it more likely that
the other subunits will bind substrate. This explains the cooperativity of substrate
binding.

When the conformation of the protein changes, the affinity of its substrate binding
sites also changes. The concerted model was extended to include the binding of al-
losteric effectors and it can be simplified by assuming that the substrate binds only to
the R conformation and the allosteric effectors bind preferentially to one of the confor-
mations — inhibitors bind only to subunits in the T conformation and activators bind
only to subunits in the R conformation. The concerted model is based on the observed
structural symmetry of regulatory enzymes. It suggests that all subunits of a given pro-
tein molecule have the same conformation, either all R or all T.

When the enzyme shifts from one conformation to the other, all subunits change
conformation in a concerted manner. Experimental data obtained with a number of en-
zymes can be explained by this simple theory. For example, many of the properties of
phosphofructokinase- 1 from E. coli fit the concerted theory. In most cases, however, the
concerted theory does not adequately account for all of the observations concerning a
particular enzyme. Their behavior is more complex than that suggested by this simple
all-or-nothing model.

The sequential model was first proposed by Daniel Koshland, George Nemethy, and
David Filmer (KNF model). It is a more general model because it allows for both
subunits to exist in two different conformations within the same multimeric protein.
The specific induced- fit version or the model is based on the idea that a ligand may in-
duce a change in the tertiary structure of each subunit to which it binds. This subunit-ligand

158

CHAPTER 5 Properties of Enzymes

complex may change the conformations of neighboring subunits to varying extents.
Like the concerted model, the sequential model assumes that only one shape has a high
affinity for the ligand but it differs from the concerted model in allowing for the exis-
tence of both high- and low- affinity subunits in a multisubunit protein (Figure 5.21b).

Hundreds of allosteric proteins have been studied and the majority show coopera-
tive binding of substrates and/or effector molecules. It has proven to be very difficult to
distinguish between the concerted and sequential models. Many proteins exhibit bind-
ing behavior that can best be explained as a mixture of the all-or-nothing shift of the
concerted model and the stepwise shift of the sequential model.

D. Regulation by Covalent Modification

▲ Figure 5.22

Regulation of mammalian pyruvate dehydroge-
nase. Pyruvate dehydrogenase, an intercon-
vertible enzyme, is inactivated by
phosphorylation catalyzed by pyruvate
dehydrogenase kinase. It is reactivated by
hydrolysis of its phosphoserine residue,
catalyzed by an allosteric hydrolase called
pyruvate dehydrogenase phosphatase.

The activity of an enzyme can be modified by the covalent attachment and removal of
groups on the polypeptide chain. Regulation by covalent modification is usually slower
than the allosteric regulation described above. It’s important to note that the covalent
modification of regulated enzymes must be reversible, otherwise it wouldn’t be a form
of regulation. The modifications usually require additional modifying enzymes for acti-
vation and inactivation. The activities of these modifying enzymes may themselves be
allosterically regulated or regulated by covalent modification. Enzymes controlled by
covalent modification are believed to generally undergo R v T transitions but they
may be frozen in one conformation or the other by a covalent substitution.

The most common type of covalent modification is phosphorylation of one or
more specific serine residues, although in some cases threonine, tyrosine, or histidine
residues are phosphorylated. An enzyme called a protein kinase catalyzes the transfer of
the terminal phosphoryl group from ATP to the appropriate serine residue of the regu-
lated enzyme. The phosphoserine of the regulated enzyme is hydrolyzed by the activity
of a protein phosphatase, releasing phosphate and returning the enzyme to its dephos-
phorylated state. Individual enzymes differ as to whether it is their phosphorylated or
dephosphorylated forms that are active.

The reactions involved in the regulation of mammalian pyruvate dehydrogenase by
covalent modification are shown in Figure 5.22. Pyruvate dehydrogenase catalyzes a re-
action that connects the pathway of glycolysis to the citric acid cycle. Phosphorylation
of pyruvate dehydrogenase, catalyzed by the allosteric enzyme pyruvate dehydrogenase
kinase, inactivates the dehydrogenase. The kinase can be activated by any of several
metabolites. Phosphorylated pyruvate dehydrogenase is reactivated under different
metabolic conditions by hydrolysis of its phosphoserine residue, catalyzed by pyruvate
dehydrogenase phosphatase.

5.10 Multienzyme Complexes and
Multifunctional Enzymes

In some cases, different enzymes that catalyze sequential reactions in the same pathway
are bound together in a multienzyme complex. In other cases, different activities may be
found on a single multifunctional polypeptide chain. The presence of multiple activities
on a single polypeptide chain is usually the result of a gene fusion event.

Some multienzyme complexes are quite stable. We will encounter several of these
complexes in other chapters. In other multienzyme complexes the proteins may be
associated more weakly (Section 4.9). Because these complexes dissociate easily it has
been difficult to demonstrate their existence and importance. Attachment to mem-
branes or cytoskeletal components is another way that enzymes may be associated.

The metabolic advantages of multienzyme complexes and multifunctional en-
zymes include the possibility of metabolite channeling. Channeling of reactants between
active sites can occur when the product of one reaction is transferred directly to the
next active site without entering the bulk solvent. This can vastly increase the rate of a
reaction by decreasing transit times for intermediates between enzymes and by produc-
ing local high concentrations of intermediates. Channeling can also protect chemically
labile intermediates from degradation by the solvent. Metabolic channeling is one way
in which enzymes can effectively couple separate reactions.

Problems 159

One of the best- characterized examples of channeling involves the enzyme trypto-
phan synthase that catalyzes the last two steps in the biosynthesis of tryptophan (Sec-
tion 17.3F). Tryptophan synthase has a tunnel that conducts a reactant between its two
active sites. The structure of the enzyme not only prevents the loss of the reactant to the
bulk solvent but also provides allosteric control to keep the reactions occurring at the
two active sites in phase.

Several other enzymes have two or three active sites connected by a molecular tun-
nel. Another mechanism for metabolite channeling involves guiding the reactant along
a path of basic amino acid side chains on the surface of coupled enzymes. The metabo-
lites (most of which are negatively charged) are directed between active sites by the elec-
trostatically positive surface path. The fatty acid synthase complex catalyzes a sequence
of seven reactions required for the synthesis of fatty acids. The structure of this complex
is described in Chapter 16 (Section 16.1).

The search for enzyme complexes and the evaluation of their catalytic and regula-
tory roles is an extremely active area of research.

The regulation of pyruvate dehydroge-
nase activity is explained in Section
13.5. An example of a signal transduc-
tion pathway involving covalent modifi-
cation is described in Section 12.6.

Summary

1. Enzymes, the catalysts of living organisms, are remarkable for
their catalytic efficiency and their substrate and reaction speci-
ficity. With few exceptions, enzymes are proteins or proteins plus
cofactors. Enzymes are grouped into six classes (oxidoreductases,
transferases, hydrolases, lyases, isomerases, and ligases) according
to the nature of the reactions they catalyze.

2. The kinetics of a chemical reaction can be described by a rate
equation.

3. Enzymes and substrates form noncovalent enzyme-substrate
complexes. Consequently, enzymatic reactions are characteris-
tically first order with respect to enzyme concentration and
typically show hyperbolic dependence on substrate concentra-
tion. The hyperbola is described by the Michaelis-Menten
equation.

4. Maximum velocity (Vmax) is reached when the substrate concen-
tration is saturating. The Michaelis constant (K m ) is equal to the
substrate concentration at half-maximal reaction velocity — that
is, at half- saturation of E with S.

5. The catalytic constant (fc cat ), or turnover number, for an enzyme
is the maximum number of molecules of substrate that can be
transformed into product per molecule of enzyme (or per active
site) per second. The ratio k cat /K m is an apparent second-order

rate constant that governs the reaction of an enzyme when the
substrate is dilute and nonsaturating. k cat /K m provides a measure
of the catalytic efficiency of an enzyme.

6. K m and V max can be obtained from plots of initial velocity at a series
of substrate concentrations and at a fixed enzyme concentration.

7. Multisubstrate reactions may follow a sequential mechanism with
binding and release events being ordered or random, or a ping-
pong mechanism.

8. Inhibitors decrease the rates of enzyme- catalyzed reactions. Re-
versible inhibitors may be competitive (increasing the apparent
value of K m without changing V max ), uncompetitive (appearing
to decrease K m and V max proportionally), noncompetitive
(appearing to decrease V max without changing K m ), or mixed.
Irreversible enzyme inhibitors form covalent bonds with the
enzyme.

9. Allosteric modulators bind to enzymes at a site other than the ac-
tive site and alter enzyme activity. Two models, the concerted
model and the sequential model, describe the cooperativity of al-
losteric enzymes. Covalent modification, usually phosphorylation,
of certain regulatory enzymes can also regulate enzyme activity.

Multienzyme complexes and multifunctional enzymes are very
common. They can channel metabolites between active sites.

Problems

1. Initial velocities have been measured for the reaction of a-chy-
motrypsin with tyrosine benzyl ester [S] at six different substrate
concentrations. Use the data below to make a reasonable estimate
of the V max and K m value for this substrate.

mM[S] 0.00125 0.01 0.04 0.10 2.0 10

(mM/min) 14 35 56 66 69 70

2. Why is the k cat /K m value used to measure the catalytic proficiency
of an enzyme?

(a) What are the upper limits for k cat /K m values for enzymes?

(b) Enzymes with k CSit /K m values approaching these upper limits
are said to have reached “catalytic perfection.” Explain.

3. Carbonic anhydrase (CA) has a 25,000-fold higher activity (fc cat =
10 6 s -1 ) than orotidine monophosphate decarboxylase (OMPD)
(fccat = 40 s -1 ). However, OMPD provides more than a 10 10 higher
“rate acceleration” than CA (Table 5.2). Explain how this is possible.

4. An enzyme that follows Michaelis-Menten kinetics has a K m of
1 ^M. The initial velocity is 0.1 ^M min -1 at a substrate concen-
tration of 100 jdM. What is the initial velocity when [S] is equal to
(a) 1 mM, (b) 1 ^M, or (c) 2 ^M?

5. Human immunodeficiency virus 1 (HIV-1) encodes a protease
(M r 21,500) that is essential for the assembly and maturation of
the virus. The protease catalyzes the hydrolysis of a heptapeptide
substrate with a /c cat of 1000 s -1 and a K m of 0.075 M.

160 CHAPTER 5 Properties of Enzymes

(a) Calculate V max for substrate hydrolysis when HIV- 1 protease
is present at 0.2 mg ml -1 .

(b) When — C(0)NH — of the heptapeptide is replaced by
— CH 2 NH — , the resulting derivative cannot be cleaved by
HIV- 1 protease and acts as an inhibitor. Under the same ex-
perimental conditions as in part (a), but in the presence of
2.5 n M inhibitor, V max is 9.3 x 10 -3 M s -1 . What kind of inhi-
bition is occurring? Is this type of inhibition expected for a
molecule of this structure?

6. Draw a graph of v 0 versus [S] for a typical enzyme reaction (a) in
the absence of an inhibitor, (b) in the presence of a competitive
inhibitor, and (c) in the presence of a noncompetitive inhibitor.

7. Sulfonamides (sulfa drugs) such as sulfanilamide are antibacterial
drugs that inhibit the enzyme dihydropteroate synthase (DS) that
is required for the synthesis of folic acid in bacteria. There is no
corresponding enzyme inhibition in animals because folic acid is
a required vitamin and cannot be synthesized. If p aminobenzoic
acid (PABA) is a substrate for DS, what type of inhibition can be
predicted for the bacterial synthase enzyme in the presence of sul-
fonamides? Draw a double reciprocal plot for this type of inhibi-
tion with correctly labeled axes and identify the uninhibited and
inhibited lines.

O O

Sulfonamides p- Aminobenzoic acid

(R = H, sulfanilamide)

8. (a) Fumarase is an enzyme in the citric acid cycle that catalyzes
the conversion of fumarate to L-malate. Given the fumarate
(substrate) concentrations and initial velocities below,
construct a Lineweaver-Burk plot and determine the V max

and K m values for the fumarase-catalyzed reaction.

Fumarate (mM)

Rate (mmol 1 1 min

02.0

2.5

03.3

3.1

05.0

3.6

10.0

4.2

(b) Fumarase has a molecular weight of 194,000 and is composed of
four identical subunits, each with an active site. If the enzyme
concentration is 1 X 10~ 2 M for the experiment in part (a),
calculate the k cat value for the reaction of fumarase with
fumarate. Note : The units for k cat are reciprocal seconds (s -1 ).

9. Covalent enzyme regulation plays an important role in the
metabolism of muscle glycogen, an energy storage molecule. The
active phosphorylated form of glycogen phosphorylase (GP) cat-
alyzes the degradation of glycogen to glucose 1 -phosphate. Using
pyruvate dehydrogenase as a model (Figure 5.23), fill in the boxes
below for the activation and inactivation of muscle glycogen
phosphorylase.

10 . Regulatory enzymes in metabolic pathways are often found at the
first step that is unique to that pathway. How does regulation at
this point improve metabolic efficiency?

11. ATCase is a regulatory enzyme at the beginning of the pathway
for the biosynthesis of pyrimidine nucleotides. ATCase exhibits
positive cooperativity and is activated in vitro by ATP and inhib-
ited by the pyrimidine nucleotide cytidine triphosphate (CTP).
Both ATP and CTP affect the K m for the substrate aspartate but
not V max . In the absence of ATP or CTP, the concentration of as-
partate required for half-maximal velocity is about 5 mM at satu-
rating concentrations of the second substrate, carbamoyl phos-
phate. Draw a v 0 versus [aspartate] plot for ATCase, and indicate
how CTP and ATP affect v 0 when [aspartate] = 5 mM.

12. The cytochrome P450 family of monooxygenase enzymes are in-
volved in the clearance of foreign compounds (including drugs)
from our body. P450s are found in many tissues, including the
liver, intestine, nasal tissues, and lung. For every drug that is ap-
proved for human use the pharmaceutical company must investi-
gate the metabolism of the drug by cytochrome P450. Many of the
adverse drug-drug interactions known to occur are a result of inter-
actions with the cytochrome P450 enzymes. A significant portion of
drugs are metabolized by one of the P450 enzymes, P450 3A4.
Human intestinal P450 3A4 is known to metabolize midazolam, a
sedative, to a hydroxylated product, U-hydroxymidazolam. The ki-
netic data given below are for the reaction catalyzed by P450 3A4.

(a) Focusing on the first two columns, determine the K m and
Umax for the enzyme using a Lineweaver-Burk plot.

(b) Ketoconazole, an antifungal, is known to cause adverse
drug-drug interactions when administered with midazolam.
Using the data in the table, determine the type of inhibition
that ketoconazole exerts on the P450-catalyzed hydroxyla-
tion of midazolam.

Rate of product
formation in the

Rate of product presence of 0.1 pM
formation ketoconazole

Midazolam(^M) (pmol 1 1 min -1 ) (pmol 1 1 min 1 )

100

156

222

323

[Adapted from Gibbs, M. A., Thummel, K. E., Shen, D. D., and
Kunze, K. L. DrugMetab. Dispos. (1999). 27:180-187]

Selected Readings 161

13. Patients who are taking certain medications are warned by their
physicians to avoid taking these medications with grapefruit
juice, which contains many compounds including bergamottin.
Cytochrome P450 3A4 is a monooxygenase that is known to me-
tabolize drugs to their inactive forms. The following results were
obtained when P450 3A4 activity was measured in the absence or
presence of bergamottin.

Bergamottin (^M)

(a) What is the effect of adding bergamottin to the P450-cat-
alyzed reaction?

(b) Why could it be dangerous for a patient to take certain
medications with grapefruit juice?

[Adapted from Wen, Y. H., Sahi, J., Urda, E., Kalkarni, S.,
Rose, K., Zheng, X., Sinclair, J. F., Cai, H., Strom, S. C., and
Kostrubsky, V. E. Drug Metab. Dispos. (2002). 30:977-984.]

14 . Use the Michaelis-Menten equation (Equation 5.14) to
demonstrate the following:

(a) v 0 becomes independent of [S] when [S]»X m .

(b) The reaction is first order with respect to S when [S] «K m .

Selected Readings

Enzyme Catalysis

Fersht, A. (1985). Enzyme Structure and Mecha-
nism , 2nd ed. (New York: W. H. Freeman).

Lewis, C. A., and Wolfenden, R. (2008). Uropor-
phyrinogen decarboxylation as a benchmark for
the catalytic proficiency of enzymes. Proc. Natl
Acad. Sci. (USA). 105:17328-17333.

Miller, B. G., and Wolfenden, R. (2002). Catalytic
proficiency: the unusual case of OMP decarboxy-
lase. Annu. Rev. Biochem. 71, 847-885.

Sigman, D. S., and Boyer, P. D., eds. (1990-1992).
The Enzymes , Vols. 19 and 20, 3rd ed. (San Diego:
Academic Press).

Webb, E. C., ed. (1992). Enzyme Nomenclature
1992: Recommendations of the Nomenclature Com-
mittee of the International Union of Biochemistry
and Molecular Biology on the Nomenclature and
Classification of Enzymes (San Diego; Academic
Press).

Enzyme Kinetics and Inhibition

Bugg, C. E., Carson, W. M., and Montgomery, J. A.
(1993). Drugs by design. Sci. Am. 269(6):92-98.

Chandrasekhar, S. (2002). Thermodynamic analy-
sis of enzyme catalysed reactions: new insights
into the Michaelis-Menten equation. Res. Cehm.
Intermed. 28:265-2 75.

Cleland, W. W. (1970). Steady State Kinetics. The
Enzymes , Vol. 2, 3rd ed., P. D. Boyer, ed. (New York:
Academic Press), pp. 1-65.

Cornish- Bowden, A. (1999). Enzyme kinetics from
a metabolic perspective. Biochem. Soc. Trans.
27:281-284.

Northrop, D. B. (1998). On the meaning of K m
and V/K in enzyme Kinetics. /. Chem. Ed.
75:1153-1157.

Radzicka, A., and Wolfenden, R. (1995). A profi-
cient enzyme. Science 267:90-93.

Segel, I. H. (1975) Enzyme Kinetics: Behavior and
Analysis of Rapid Equilibrium and Steady State
Enzyme Systems (New York: Wiley-Interscience).

Regulated Enzymes

Ackers, G. K., Doyle, M. L., Myers, D., and Daugh-
erty, M. A. (1992). Molecular code for cooperativ-
ity in hemoglobin. Science 255:54-63.

Barford, D. (1991). Molecular mechanisms for the
control of enzymic activity by protein phosphory-
lation. Biochim. Biophys. Acta 1133:55-62.

Hilser, V. J. (2010). An ensemble view of allostery.
Science 327:653-654.

Hurley, J. H., Dean, A. M., Sohl, J. L., Koshland, D.
E., Jr., and Stroud, R. M. (1990). Regulation of an
enzyme by phosphorylation at the active site.
Science 249:1012-1016.

Schirmer, T., and Evans, P. R. (1990). Structural
basis of the allosteric behavior of phosphofructok-
inase. Nature 343:140-145.

Metabolite Channeling

Pan, P., Woehl, E., and Dunn, M. F. (1997). Protein
architecture, dynamics and allostery in tryptophan
synthase channeling. Trends Biochem. Sci.

22:22-27.

Velot, C., Mixon, M. B., Teige, M., and Srere, P. A.
(1997). Model of a quinary structure between
Krebs TCA cycle enzymes: a model for the
metabolon. Biochemistry 36:14271-14276.

Mechanisms of Enzymes

T he previous chapter described some general properties of enzymes with an
emphasis on enzyme kinetics. In this chapter, we see how enzymes catalyze reactions
by studying the molecular details of catalyzed reactions. Individual enzyme
mechanisms have been deduced by a variety of methods including kinetic experiments,
protein structural studies, and studies of nonenzymatic model reactions. The results of
such studies show that the extraordinary catalytic ability of enzymes results from simple
physical and chemical properties, especially the binding and proper positioning of reac-
tants in the active sites of enzymes. Chemistry, physics, and biochemistry have combined
to take much of the mystery out of enzymes and recombinant DNA technology now
allows us to test the theories proposed by enzyme chemists. Observations for which
there were no explanations just a half-century ago are now thoroughly understood.

The mechanisms of many enzymes are well established and they give us a general pic-
ture of how enzymes function as catalysts. We begin this chapter with a review of simple
chemical mechanisms, followed by a brief discussion of catalysis. We then examine the
major modes of enzymatic catalysis: acid-base and covalent catalysis (classified as chemi-
cal effects) and substrate binding and transition state stabilization (classified as binding
effects). We end the chapter with some specific examples of enzyme mechanisms.

I think that enzymes are molecules
that are complementary in structure
to the activated complexes of the
reactions that they catalyze.

—Linus Pauling (1948)

6.1 The Terminology of Mechanistic Chemistry

The mechanism of a reaction is a detailed description of the molecular, atomic, and
even subatomic events that occur during the reaction. Reactants, products, and any in-
termediates must be identified. A number of laboratory techniques are used to deter-
mine the mechanism of a reaction. For example, the use of isotopically labeled reactants
can trace the path of individual atoms and kinetic techniques can measure the changes in
chemical bonds of a reactant or solvent during the reaction. Study of the stereochemical
changes that occur during the reaction can give a three-dimensional view of the process.
For any proposed enzyme mechanism, the mechanistic information about the reactants
and intermediates must be coordinated with the three-dimensional structure of the en-
zyme. This is an important part of understanding structure-function relationships —
one of the main themes in biochemistry.

Top: A step from the mechanism of the triose phosphate isomerase reaction.

162

6.1 The Terminology of Mechanistic Chemistry

163

Enzymatic mechanisms are described using the same symbolism developed in or-
ganic chemistry to represent the breaking and forming of chemical bonds. The move-
ment of electrons is the key to understanding chemical (and enzymatic) reactions. We
will review chemical mechanisms in this section and in the following sections we will
discuss catalysis and present several specific enzyme mechanisms. This discussion
should provide sufficient background for you to understand all the enzyme -catalyzed
reactions presented in this book.

A. Nucleophilic Substitutions

Many chemical reactions have ionic substrate, intermediates, or products. There are two
types of ionic molecules: one species is electron rich, or nucleophilic, and the other species
is electron poor, or electrophilic (Section 2.6). A nucleophile has a negative charge or an
unshared electron pair. We usually think of the nucleophile as attacking the electrophile
and call the mechanism a nucleophilic attack or a nucleophilic substitution. In mechanistic
chemistry, the movement of a pair of electrons is represented by a curved arrow pointing
from the available electrons of the nucleophile to the electrophilic center. These “electron
pushing” diagrams depict the breaking of an existing covalent bond or the formation of a
new covalent bond. The reaction mechanism usually involves an intermediate.

Many biochemical reactions are group transfer reactions where a group is moved
from one molecule to another. Many of these reactions involve a charged intermediate.
The transfer of an acyl group, for example, can be written as the general mechanism

x 0

( 6 . 1 )

The nucleophile Y® attacks the carbonyl carbon (i.e., adds to the carbonyl carbon atom)
to form a tetrahedral addition intermediate from which is eliminated. is called
the leaving group — the group displaced by the attacking nucleophile. This is an example

of a nucleophilic substitution reaction.

Another type of nucleophilic substitution involves direct displacement. In this
mechanism, the attacking group, or molecule, adds to the face of the central atom op-
posite the leaving group to form a transition state having five groups associated with the
central atom. This transition state is unstable. It has a structure between that of the re-
actant and that of the product. (Transition states are shown in square brackets to
identify them as unstable, transient entities.)

i \j

L r 3 J

Transition state

R 2 R,

\ /

/ \

X Rq

+ Y

( 6 . 2 )

Note that both types of nucleophilic substitution mechanisms involve a transitory
state. In the first type (Reaction 6.1), the reaction proceeds in a stepwise manner form-
ing an intermediate molecule that may be stable enough to be detected. In the second
type of mechanism (Reaction 6.2), the addition of the attacking nucleophile and the
displacement of the leaving group occur simultaneously. The transition state is not a
stable intermediate.

B. Cleavage Reactions

We will also encounter cleavage reactions. Covalent bonds can be cleaved in two ways: ei-
ther both electrons can stay with one atom or one electron can remain with each atom.

Transition states are discussed further
in Section 6.2.

164

CHAPTER 6 Mechanisms of Enzymes

The two electrons will stay with one atom in most reactions so that an ionic intermediate
and a leaving group are formed. For example, cleavage of a C — H bond almost always
produces two ions. If the carbon atom retains both electrons then the carbon- containing
compound becomes a carbanion and the other product is a proton.

R 3 — c— H > R 3 — O e + H©

Carbanion Proton (6-3)

If the carbon atom loses both electrons, the carbon-containing compound becomes a
cationic ion called a carbocation and the hydride ion carries a pair of electrons.

R 3 — c — H ■* R 3 — C© + H©

Carbocation Hydride ^

In the second, less common, type of bond cleavage, one electron remains with each
product to form two free radicals that are usually very unstable. (A free radical, or radi-
cal, is a molecule or atom with an unpaired electron.)

RtO — OR 2 > RiO + -OR 2 (6.5)

Loss of Electrons = Oxidation (LEO)
Gain of Electrons = Reduction (GER)

Remember the phrase: LEO (the lion)
says GER

Oxidation is Loss (OIL)

Reduction is Gain (RIG)

Remember the phrase: OIL RIG

C. Oxidation-Reduction Reactions

Oxidation-reduction reactions are central to the supply of biological energy. In an
oxidation-reduction (redox) reaction, electrons from one molecule are transferred to
another. The terminology here can be a bit confusing so it’s important to master the
meaning of the words oxidation and reduction — they will come up repeatedly in the rest
of the book. Oxidation is the loss of electrons: a substance that is oxidized will have fewer
electrons when the reaction is complete. Reduction is the gain of electrons: a substance
that gains electrons in a reaction is reduced. Oxidation and reduction reactions always
occur together. One substrate is oxidized and the other is reduced. An oxidizing agent is
a substance that causes an oxidation — it takes electrons from the substrate that is oxi-
dized. Thus, oxidizing agents gain electrons (i.e., they are reduced). A reducing agent is
a substance that donates electrons (and is oxidized in the process).

Oxidations can take several forms, such as removal of hydrogen (dehydrogena-
tion), addition of oxygen, or removal of electrons. Dehydrogenation is the most com-
mon form of biological oxidation. Recall that oxidoreductases (enzymes that catalyze
oxidation-reduction reactions) represent a large class of enzymes and dehydrogenases
(enzymes that catalyze removal of hydrogen) are a major subclass of oxidoreductases
(Section 5.1).

Most dehydrogenations occur by C — H bond cleavage producing a hydride ion
(H®). The substrate is oxidized because it loses the electrons associated with the
hydride ion. Such reactions will be accompanied by a corresponding reduction where
another substrate gains electrons by reacting with the hydride ion. The dehydrogena-
tion of lactate (Equation 5.1) is an example of the removal of hydrogen. In this case, the
oxidation of lactate is coupled to the reduction of the coenzyme NAD®. The role of
cofactors in oxidation-reduction reactions will be discussed in the next chapter
(Section 7.3) and the free energy of these reactions is described in Section 10.9.

6.2 Catalysts Stabilize Transition States

In order to understand catalysis it’s necessary to appreciate the importance of transition
states and intermediates in chemical reactions. The rate of a chemical reaction depends
on how often reacting molecules collide in such a way that a reaction is favored. The col-
liding substances must be in the correct orientation and must possess sufficient energy to
approach the physical configuration of the atoms and bonds of the final product.

As mentioned above, the transition state is an unstable arrangement of atoms in
which chemical bonds are in the process of being formed or broken. Transition states

6.2 Catalysts Stabilize Transition States 165

◄ Figure 6.1

Energy diagram for a single-step reaction. The

upper arrow shows the activation energy for
the forward reaction. Molecules of substrate
that have more free energy than the activa-
tion energy pass over the activation barrier
and become molecules of product. For reac-
tions with a high activation barrier, energy in
the form of heat must be provided in order
for the reaction to proceed.

Course of the reaction >

(Reaction coordinate)

have extremely short lifetimes of about 10 -14 to 10 -13 second, the time of one bond vi-
bration. Although they are very difficult to detect, their structures can be predicted. The
energy required to reach the transition state from the ground state of the reactants is called
the activation energy of the reaction and is often referred to as the activation barrier.

The progress of a reaction can be represented by an energy diagram, or energy pro-
file. Figure 6.1 is an example that shows the conversion of a substrate (reactant) to a
product in a single step. The y axis shows the free energies of the reacting species. The
x axis, called the reaction coordinate , measures the progress of the reaction, beginning
with the substrate on the left and proceeding to the product on the right. This axis is not
time but rather the progress of bond breaking and bond formation of a particular mol-
ecule. The transition state occurs at the peak of the activation barrier — this is the energy
level that must be exceeded for the reaction to proceed. The lower the barrier the more
stable the transition state and the more often the reaction proceeds.

Intermediates, unlike transition states, can be sufficiently stable to be detected or iso-
lated. When there is an intermediate in a reaction, the energy diagram has a trough that
represents the free energy of the intermediate as shown in Figure 6.2. This reaction has two
transition states, one preceding formation of the intermediate and one preceding its con-
version to product. The slowest step, the rate- determining or rate-limiting step, is the step
with the highest energy transition state. In Figure 6.2, the rate-determining step is the for-
mation of the intermediate. The intermediate is metastable because relatively little energy is
required for the intermediate either to continue to product or to revert to the original reac-
tant. Proposed intermediates that are too short-lived to be isolated or detected are often en-
closed in square brackets like transition states, which they presumably closely resemble.

Catalysts create reaction pathways that have lower activation energies than those of
uncatalyzed reactions. Catalysts participate directly in reactions by stabilizing the tran-
sition states along the reaction pathways. Enzymes are catalysts that accelerate reactions
by lowering the overall activation energy. They achieve rate enhancement by providing
a multistep pathway (with one or several intermediates) in which each of the steps has
lower activation energy than the corresponding stages in the nonenzymatic reaction.

The first step in an enzymatic reaction is the formation of a noncovalent
enzyme-substrate complex, ES. In a reaction between A and B, formation of the EAB
complex collects and positions the reactants making the probability of reaction much
higher for the enzyme -catalyzed reaction than for the uncatalyzed reaction. Figures 6.3a
and 6.3b show a hypothetical case in which substrate binding is the only mode of
catalysis by an enzyme. In this example, the activation energy is lowered by bringing the
reactants together in the substrate binding site. Correct substrate binding accounts for a
large part of the catalytic power of enzymes.

The active sites of enzymes bind substrates and products. They also bind transition
states. In fact, transition states are likely to bind to active sites much more tightly than

KEY CONCEPT

Transition states are unstable molecules
with free energies higher than either the
substrate or the product.

The meaning of activation energy is
described in Section 1.4D.

▲ Figure 6.2

Energy diagram for a reaction with an interme-
diate. The intermediate occurs in the trough
between the two transition states. The rate-
determining step in the forward direction is
formation of the first transition state, the
step with the higher energy transition state.
S represents the substrate, and P represents
the product.

166 CHAPTER 6 Mechanisms of Enzymes

(a) Uncatalyzed reaction (b) Effect of reactants being bound

by enzyme

▲ Figure 6.3

Enzymatic catalysis of the reaction A + B — * A — B. (a) Energy diagram for an uncatalyzed reaction, (b) Effect of reactant binding. Collection of the two
reactants in the EAB complex properly positions them for reaction, makes formation of the transition state more frequent, and hence lowers the
activation energy, (c) Effect of transition-state stabilization. An enzyme binds the transition state more tightly than it binds substrates, further lower-
ing the activation energy. Thus, an enzymatic reaction has a much lower activation energy than an uncatalyzed reaction. (The breaks in the reaction
curves indicate that the enzymes provide multistep pathways.)

substrates do. The extra binding interactions stabilize the transition state, further lowering
the activation energy (Figure 6.3c). We will see that the binding of substrates followed by
the binding of transition states provides the greatest rate acceleration in enzyme catalysis.

We return to binding phenomena later in this chapter after we examine the chemi-
cal processes that underlie enzyme function. (Note that enzyme -catalyzed reactions are
usually reversible. The same principles apply to the reverse reaction. The activation en-
ergy is lowered by binding the “products” and stabilizing the transition state.)

In addition to reactive amino acid
residues, there may be metal ions or
coenzymes in the active site. The role
of these cofactors in enzyme catalysis
is described in Chapter 7.

6.3 Chemical Modes of Enzymatic Catalysis

The formation of an ES complex places reactants in proximity to reactive amino acid
residues in the enzyme active site. Ionizable side chains participate in two kinds of
chemical catalysis; acid-base catalysis and covalent catalysis. These are the two major
chemical modes of catalysis.

A. Polar Amino Acid Residues in Active Sites

The active site cavity of an enzyme is generally lined with hydrophobic amino acid
residues. However, a few polar, ionizable residues (and a few molecules of water) may
also be present in the active site. Polar amino acid residues (or sometimes coenzymes)
undergo chemical changes during enzymatic catalysis. These residues make up much of
the catalytic center of the enzyme.

Table 6.1 lists the ionizable residues found in the active sites of enzymes. Histidine,
which has a p K a of about 6 to 7 in proteins, is often an acceptor or a donor of protons.
Aspartate, glutamate, and occasionally lysine can also participate in proton transfer.
Certain amino acids, such as serine and cysteine, are commonly involved in group-
transfer reactions. At neutral pH, aspartate and glutamate usually have negative charges,
and lysine and arginine have positive charges. These anions and cations can serve as
sites for electrostatic binding of oppositely charged groups on substrates.

6.3 Chemical Modes of Enzymatic Catalysis 167

BOX 6.1 SITE-DIRECTED MUTAGENESIS MODIFIES ENZYMES

It is possible to test the functions of the amino acid side
chains of an enzyme using the technique of site-directed mu-
tagenesis (see Section 23.10). This technique has had a huge
impact on our understanding of structure-function relation-
ships of enzymes.

In site-directed mutagenesis, a desired mutation is engi-
neered directly into a gene by synthesizing an oligonucleotide
that contains the mutation flanked by sequences identical to
the target gene. When this oligonucleotide is used as a primer
for DNA replication in vitro , the new copy of the gene contains
the desired mutation. Since alterations can be made at any
position in a gene, specific changes in proteins can be engineered
allowing direct testing of hypotheses about the functional
role of key amino acid residues. Site-directed mutagenesis is

commonly used to introduce single codon mutations into
genes, resulting in single amino acid substitutions.

The mutated gene can be introduced into bacterial cells
where modified enzymes are synthesized from the gene. The
structure and activity of the mutant protein can then be ana-
lyzed to see the effect of changing an individual amino acid.

■

i jmM

'ff iPfjr

i A '

'v, *.•

f NO

Ordinary

'MIKE

Michael Smith h Nobel Laureate

Eric Darner e- Carol i no Astell

▲ Michael Smith (1932-2000), received
the Nobel Prize in Chemistry in 1993 for
inventing site-directed mutagenesis.

Single-stranded vector
containing sequence to
be altered

Hybridization

Extension

Ligation

Three-base

mismatch

◄ Oligonucleotide-directed, site-specific
mutagenesis. A synthetic oligonucleotide
containing the desired change (3 bp) is
annealed to the single-stranded vector
containing the sequence to be altered. The
synthetic oligonucleotide serves as a primer
for the synthesis of a complementary strand.
The double-stranded, circular heteroduplex
is transformed into E. coli cells where repli-
cation produces mutant and wild-type DNA
molecules.

Transform cells

Mutant

Replication

Wild type

168 CHAPTER 6 Mechanisms of Enzymes

Table 6.1 Catalytic functions of reactive groups of ionizable amino acids

Amino acid

Reactive

group

Net charge
at pH 7

Principal functions

Aspartate

—coo©

-1

Cation binding; proton transfer

Glutamate

—coo©

-1

Cation binding; proton transfer

Histidine

Imidazole

Near 0

Proton transfer

Cysteine

— CH 2 SH

Near 0

Covalent binding of acyl groups

Tyrosine

Phenol

Hydrogen bonding to ligands

Lysine

NH^

+ 1

Anion binding; proton transfer

Arginine

Guanidinium

+ 1

Anion binding

Serine

— CH 2 OH

Covalent binding of acyl groups

Table 6.2 Typical p K a values of ionizable
groups of amino acids in proteins

Group

P*a

Terminal a-carboxyl

3-4

Side-chain carboxyl

4-5

Imidazole

6-7

Terminal a-amino

7.5-9

Thiol

8-9.5

Phenol

9.5-10

e-Amino

-10

Guanidine

-12

Hydroxymethyl

-16

Table 6.3 Frequency distribution of

catalytic residues in enzymes

% of catalytic
residues

% of all
residues

His

Asp

Arg

Glu

Lys

Cys

Tyr

Asn

Ser

Gly

The piC a values of the ionizable groups of amino acid residues in proteins may dif-
fer from the values of the same groups in free amino acids (Section 3.4). Table 6.2 lists
the typical p K a values of ionizable groups of amino acid residues in proteins. Compare
these ranges to the exact values for free amino acids in Table 3.2. A given ionizable
group can have different p K a values within a protein because of differing microenviron-
ments. These differences are usually small but can be significant.

Occasionally, the side chain of a catalytic amino acid residue exhibits a p K a quite
different from the one shown in Table 6.2. Bearing in mind that p K a values may be per-
turbed, one can test whether particular amino acids participate in a reaction by exam-
ining the effect of pH on the reaction rate. If the change in rate correlates with the p K a
of a certain ionic amino acid (Section 6. 3D), a residue of that amino acid may take
part in catalysis.

Only a small number of amino acid residues participate directly in catalyzing reac-
tions. Most residues contribute in an indirect way by helping to maintain the correct
three-dimensional structure of a protein. As we saw in Chapter 4, the majority of amino
acid residues are not evolutionarily conserved.

In vitro mutagenesis studies of enzymes have confirmed that most amino acid sub-
stitutions have little effect on enzyme activity. Nevertheless, every enzyme has a few key
residues that are absolutely essential for catalysis. Some of these residues are directly in-
volved in the catalytic mechanism, often by acting as an acid or base catalyst or a nucle-
ophile. Other residues act indirectly to assist or enhance the role of a key residue. Other
roles for key catalytic residues include substrate binding, stabilization of the transition
state, and interacting with essential cofactors.

Enzymes usually have between two and six key catalytic residues. The top ten cat-
alytic residues are listed in Table 6.3. The charged residues, His, Asp, Arg, Glu, and Lys
account for almost two-thirds of all catalytic residues. This makes sense since charged
side chains are more likely to act as acids, bases, and nucleophiles. They are also more
likely to play a role in binding substrates or transition states. The number one catalytic
residue is histidine. Histidine is 6 times more likely to be involved in catalysis than its
abundance in proteins would suggest.

B. Acid-Base Catalysis

In acid-base catalysis, the acceleration of a reaction is achieved by catalytic transfer of a
proton. Acid-base catalysis is the most common form of catalysis in organic chemistry
and it’s also common in enzymatic reactions. Enzymes that employ acid-base catalysis
rely on amino acid side chains that can donate and accept protons under the nearly neu-
tral pH conditions of cells. This type of acid-base catalysis, involving proton-transferring
agents, is termed general acid-base catalysis. (Catalysis by H® or OH® is termed specific
acid or specific base catalysis.) In effect, the active sites of these enzymes provide the bio-
logical equivalent of a solution of acid or base.

It is convenient to use B: to represent a base, or proton acceptor, and BH® to repre-
sent its conjugate acid, a proton donor. (This acid-base pair can also be written as

6.3 Chemical Modes of Enzymatic Catalysis

169

HA/A©.) a proton acceptor can assist reactions in two ways: (1) it can cleave
O — H, N — H, or even some C — H bonds by removing a proton

S' 'A • ©

— X^-pH :B < > — H — B (6.6)

and (2) the general base B: can participate in the cleavage of other bonds involving car-
bon, such as a C — N bond, by generating the equivalent of OH© in neutral solution
through removal of a proton from a molecule of water.

(°0

— C — N

H. ©

C B J

— c — OH +

( 6 . 7 )

The general acid BH© can also assist in bond cleavage. A covalent bond may break
more easily if one of its atoms is protonated. For example,

R © + OH© R-OH

H©

~T~

H ©

R — onf R @ + h 2 o

( 6 . 8 )

BH© catalyzes bond cleavage by donating a proton to an atom (such as the oxygen of
R — OH in Equation 6.8), thereby making bonds to that atom more labile. In all reac-
tions involving BH© the reverse reaction is catalyzed by B:, and vice versa.

Histidine is an ideal group for proton transfer at neutral pH values because the
imidazole/imidazolium of the side chain has a p X a of about 6 to 7 in most proteins. We
have seen that histidine is a common catalytic residue. In the following sections, we will
examine some specific roles of histidine side chains.

KEY CONCEPT

In acid-base catalysis, the reaction
requires specific amino acid side chains
that can donate and accept protons.

C. Covalent Catalysis

In covalent catalysis, a substrate is bound covalently to the enzyme to form a reactive in-
termediate. The reacting side chain of the enzyme can be either a nucleophile or an
electrophile. Nucleophilic catalysis is more common. In the second step of the reaction,
a portion of the substrate is transferred from the intermediate to a second substrate. For
example, the group X can be transferred from molecule A — X to molecule B in the fol-
lowing two steps via the covalent ES complex X — E:

A— X + E X — E + A

( 6 . 9 )

and

X — E + B B — X + E (6.10)

This is a common mechanism for coupling two different reactions in biochemistry.
Recall that the ability to couple reactions is one of the important properties of enzymes
(Chapter 5; “Introduction”). Transferases, one of the six classes of enzymes (Section 5.1),
catalyze group -transfer reactions in this manner and hydrolases catalyze a special kind
of group-transfer reaction where water is the acceptor. Transferases and hydrolases
together make up more than half of known enzymes.

The reaction catalyzed by bacterial sucrose phosphorylase is an example of group
transfer by covalent catalysis. (Sucrose is composed of one glucose residue and one
fructose residue.)

Sucrose + Pj Glucose 1 -phosphate + Fructose

( 6 . 11 )

170 CHAPTER 6 Mechanisms of Enzymes

Figure 6.4 ►

Covalent catalysis. The enzyme A/-acetyl-
D-neuraminic acid lyase from Escherichia
coli catalyzes the condensation of pyruvate
and A/-acetyl-D-mannosamine to form
A/-acetyl-D-neuraminic acid (see Section
8.7C). One of the intermediates in the reac-
tion is a Schiff base (see Fig. 5.15) between
pyruvate (black carbon atoms) and a lysine
reside. The intermediate is stabilized by
hydrogen bonds with other amino acid side
chains. [PDB 2WKJ]

KEY CONCEPT

In covalent catalysis mechanisms, the
enzyme participates directly in the
reaction. It reacts with a substrate and
an intermediate containing the enzyme is
produced. The reaction is not complete
until free enzyme is regenerated.

23456789 10 11

The first chemical step in the reaction is formation of a covalent glucosyl-enzyme inter-
mediate. In this case, sucrose is equivalent to A — X and glucose is equivalent to X in
Reaction 6.9.

Sucrose + Enzyme G I ucosyl- Enzyme + Fructose (6.12)

The covalent ES intermediate can donate the glucose unit either to another mole-
cule of fructose, in the reverse of Reaction 6.12, or to phosphate (which is equivalent to
B in Reaction 6.10).

Glucosyl-Enzyme + ^ = - Glucose 1 -phosphate + Enzyme (6.13)

Proof that an enzyme mechanism relies on covalent catalysis often requires the iso-
lation or detection of an intermediate and demonstration that it is sufficiently reactive.
In some cases, the covalently bound intermediate is seen in the crystal structure of an
enzyme, and this is direct proof of covalent catalysis (Figure 6.4 ).

D. pH Affects Enzymatic Rates

The effect of pH on the reaction rate of an enzyme can suggest which ionizable amino
acid residues are in its active site. Sensitivity to pH usually reflects an alteration in the
ionization state of one or more residues involved in catalysis, although occasionally substrate
binding is affected. A plot of reaction velocity versus pH most often yields a bell-shaped
curve provided the enzyme is not denatured when the pH is altered.

A good example is the pH versus rate profile for papain, a protease isolated from
papaya fruit (Figure 6.5). The bell- shaped pH profile can be explained by assuming that
the ascending portion of the curve represents the deprotonation of an active-site amino
acid residue (B) and the descending portion represents the deprotonation of a second
active-site amino acid residue (A). The two inflection points approximate the pX a values of
the two ionizable residues. A simple bell-shaped curve is the result of two overlapping

◄ Figure 6.5

pH vs rate profile for papain. The left and right segments of the bell-shaped curve represent the titra-
tions of the side chains of active-site amino acids. The inflection point at pH 4.2 reflects the p K a of
Cys-25, and the inflection point at pH 8.2 reflects the p K a of His-159. The enzyme is active only
when these ionic groups are present as the thiolate-imidazolium ion pair.

6.4 Diffusion-Controlled Reactions 171

titrations. The side chain of A (R A ) must be protonated for activity and the side chain of
B (R b ) must be unprotonated.

H® H® H®

Ra Rb

H®

Ra Rb

n —

1 1

-c a -c a -

n —

Inactive

^H®

Active

H®

Inactive

At the pH optimum, midway between the two pK a values, the greatest number of
enzyme molecules is in the active form with residue A protonated. Not all pH profiles
are bell-shaped. A pH profile is a sigmoidal curve if only one ionizable amino acid
residue participates in catalysis and it can have a more complicated shape if more than
two ionizable groups participate. Enzymes are routinely assayed near their optimal pH,
which is maintained using appropriate buffers.

The pH versus rate graph for papain has inflection points at pH 4.2 and pH 8.2,
suggesting that the activity of papain depends on two active-site amino acid residues
with p K a values of about 4 and 8. These ionizable residues are a nucleophilic cysteine
(Cys-25) and a proton-donating imidazolium group of histidine (His- 159) (Figure 6.6).
The side chain of cysteine normally has a p K a value of 8 to 9.5 but in the active site of
papain the piC a of Cys-25 is greatly perturbed to 3.4. The p K a of the His- 159 residue is
perturbed to 8.3. The inflection points on the pH profile do not correspond exactly to the
piC a values of Cys-25 and His- 159 because the ionization of additional groups contributes
slightly to the overall shape of the curve. Three ionic forms of the catalytic center of papain
are shown in Figure 6.7. The enzyme is active only when the thiolate group and the im-
idazolium group form an ion pair (as in the upper tautomer of the middle pair).

▲ Figure 6.6 Ionizable residues in papain.

Model of papain, showing bal l-and-stick
models of the active-site histidine and
cysteine side chain. The imidazole nitrogen
atoms are blue, and the sulfur atom is
yellow.

6.4 Diffusion-Controlled Reactions

A few enzymes catalyze reactions at rates approaching the upper physical limit of reac-
tions in solution. This theoretical upper limit is the rate of diffusion of reactants into
the active site. A reaction that occurs with every collision between reactant molecules is
termed a diffusion controlled reaction or a diffusion-limited reaction. Under physiological
conditions the diffusion- controlled rate is about 10 8 to 10 9 M -1 s _1 . Compare this theo-
retical maximum to the apparent second- order rate constants (k cat /K m ) for five very fast
enzymes listed in Table 6.4.

The binding of a substrate to an enzyme is a rapid reaction. If the rest of the reac-
tion is simple and fast, the binding step may be the rate-determining step and the over-
all rate of the reaction may approach the upper limit for catalysis. Only a few types of
chemical reactions can proceed this quickly. These include association reactions, some
proton transfers, and electron transfers. The reactions catalyzed by all the enzymes
listed in Table 6.4 are so simple that the rate- determining steps are roughly as fast as

Table 6.4 Enzymes with second-order rate constants near the upper limit

Enzyme

Substrate

*cat/Km(M 1 S V

Catalase

h 2 o 2

4 X 10 7

Acetylcholinesterase

Acetylcholine

2 X 10 8

Triose phosphate isomerase

D-Glyceraldehyde 3-phosphate

4 X 10 8

Fumarase

Fumarate

10 9

Superoxide dismutase

•op

2 X 10 9

*The ratio k cat /K m is the apparent second-order rate constant for the enzyme-catalyzed reaction E + S — » E + P.
For these enzymes, the formation of the ES complex can be the slowest step.

172 CHAPTER 6 Mechanisms of Enzymes

His

H®^

pK a = 3.4

binding of substrates to the enzymes. They catalyze diffusion- controlled reactions. We will
now look at two of these enzymes in detail: triose phosphate isomerase and superoxide
dismutase.

A. Triose Phosphate Isomerase

Triose phosphate isomerase catalyzes the rapid interconversion of dihydroxyacetone
phosphate (DHAP) and glycer aldehyde 3 -phosphate (G3P) in the glycolysis and gluco-
neogenesis pathways (Chapters 11 and 12).

His

Cys

~T ch 2

izC \ f=(

s — H — :N^NH

H ©^

p/C a = 8.3

His

1 CH 2 OH

2 C=0

3 ch 2 opo 3 ®

Triose

phosphate

isomerase

Dihydroxyacetone
phosphate (DHAP)

H O

H — C — OH

CH.OPO

o-Glyceraldehyde

3-phosphate

(G3P)

(6.15)

The reaction proceeds by shifting protons from the carbon atom 1 of DHAP to the
carbon atom 2 (Figure 6.8). Triose phosphate isomerase has two ionizable active-site
residues: glutamate that acts as a general acid-base catalyst, and histidine that shuttles a
proton between oxygen atoms of an enzyme-bound intermediate. When dihydroxyace-
tone phosphate (DHAP) binds, the carbonyl oxygen forms a hydrogen bond with the
imidazole group of His-95. The carboxylate group of Glu-165 removes a proton from
C-l of the substrate to form an enoldiolate transition state (Figure 6.8, top). The tran-
sition-state molecule is rapidly converted to a stable enediol intermediate (middle,
Figure 6.8). This intermediate is then converted via a second enediolate transition state
to D-glyceraldehyde 3-phosphate (G3P).

In this reaction, the proton-donating form of histidine appears to be the neutral
species and the proton-accepting species appears to be the imidazolate. The hydrogen
bonds formed between histidine and the intermediates in this mechanism appear to be
unusually strong.

▲ Figure 6.7 The activity of papain depends
on two ionizable residues, histidine (His-159)
and cysteine (Cys-25), in the active site. Three
ionic forms of these residues are shown.

Only the upper tautomer of the middle pair
is active.

NH — CH £ v/vnyvo

CH 2

J©

HN^N:

-NH — CH — C

ch 2

(6.16)

:N'P/N: Imidazolate

The imidazolate form of a histidine residue is unusual; the triose phosphate isomerase
mechanism was the first enzymatic mechanism in which this form was implicated.

The enediol intermediate is stable and in order to prevent it from diffusing out of
the active site, triose phosphate isomerase has evolved a “locking” mechanism to seal the
active site until the reaction is complete. When substrate binds, a flexible loop of the
protein moves to cover the active site and prevent release of the enediol intermediate
(Figure 6.9).

The rate constants of all four kinetically measurable enzymatic steps have been
determined.

(1) (2) (3)

E + DHAP E-DHAP E-Intermediate

(4)

E-G3P E + G3P

(6.17)

6.4 Diffusion-Controlled Reactions 173

His-95

▲ Figure 6.8

General acid-base catalysis mechanism proposed for the
reaction catalyzed by triose phosphate isomerase.

▲ Figure 6.9

Structure of yeast ( Saccharomyces cerevisiae) triose phosphate isomerase. The location of the substrate is indicated by the space-filling model of a sub-
strate analog, (a) The structure of the “open loop” form of the enzyme when the active site is unoccupied, (b) The structure when the loop has closed
over the active site to prevent release of the enediol intermediate before the reaction is completed.

174 CHAPTER 6 Mechanisms of Enzymes

Figure 6.10 ►

Energy diagram for the reaction catalyzed by
triose phosphate isomerase. [Adapted from
Raines, R. T., Sutton, E. L., Strauss, D. R.,
Gilbert, W., and Knowles, J. R. (1986).
Reaction energetics of a mutant triose
phosphate isomerase in which the active-
site glutamate has been changed to
aspartate. Biochem. 25:7142-7154.]

The energy diagram constructed from these rate constants is shown in Figure 6.10.
Note that all the barriers for the enzyme are approximately the same height. This means
that the steps are balanced, and no single step is rate-limiting. The physical step of S
binding to E is rapid but not much faster than the subsequent chemical steps in the re-
action sequence. The value of the second- order rate constant k cat /K m for the conversion
of glyceraldehyde 3 -phosphate to dihydroxyacetone phosphate is 4 X 10 8 M -1 s _1 ,
which is close to the theoretical rate of a diffusion-controlled reaction. It appears that
this isomerase has achieved its maximum possible efficiency as a catalyst.

BOX 6.2 THE “PERFECT ENZYME”?

Much of our understanding of the mechanism of triose
phosphate isomerase (TPI) comes from the lab of Jeremy
Knowles at Harvard University (Cambridge, MA, USA). He
points out that the enzyme has achieved catalytic perfection
because the overall rate of the reaction is limited only by the
rate of diffusion of substrate into the active site. TPI cant
work any faster than this!

This has led many people to declare that TPI is the
“perfect enzyme” because it has evolved to be so efficient.
However, as Knowles and his coworkers have explained, the
“perfect enzyme” isn’t necessarily one that has evolved the
maximum reaction rate. Most enzymes are not under selec-
tive pressure to increase their rate of reaction because they
are part of a metabolic pathway that meets the cell’s needs at
less than optimal rates.

Even if it would be beneficial to increase the overall flux in
a pathway (i.e., produce more of the end product per second),
an individual enzyme need only keep up with the slowest
enzyme in the pathway in order to achieve “perfection.” The
slowest enzyme might be catalyzing a very complicated reac-
tion and might be very efficient. In this case, there will be no
selective pressure on the other enzymes to evolve faster
mechanisms and they are all “perfect enzymes.”

In all species, triose phosphate isomerase is part of the
gluconeogenesis pathway leading to the synthesis of glucose.
In most species, it also plays a role in the reverse pathway
where glucose is degraded (glycolysis). The enzyme is very
ancient, and all versions — bacterial and eukaryotic — have
achieved catalytic perfection. The two enzymes on either side of
the reaction pathway, aldolase and glyceraldehyde 3 -phosphate

dehydrogenase (Section 11.2), are much slower. Thus, it is by
no means obvious why TPI works as fast as it does.

The important point to keep in mind is that the vast majority
of enzymes have not evolved catalytic perfection because their
in vivo rates are “perfectly” adequate for the needs of the cell.

▲ The Perfect Game. New York Yankees
catcher Yogi Berra congratulates Don Larson
for pitching a perfect game in the 1956
World Series against the Brooklyn Dodgers.
Perfect games are rare in baseball but there
are many “perfect enzymes.”

6.5 Modes of Enzymatic Catalysis 175

B. Superoxide Dismutase

Superoxide dismutase is an even faster catalyst than triose phosphate isomerase. Super-
oxide dismutase catalyzes the very rapid removal of the toxic superoxide radical anion,
•0 2 ®, a by-product of oxidative metabolism. The enzyme catalyzes the conversion of
superoxide to molecular oxygen and hydrogen peroxide, which is rapidly removed by
the subsequent action of enzymes such as catalase.

4 H

4-0

2 Superoxide
dismutase

2 0 2

» 2 H 2 0 2

Catalase

2 H 2 0 + 0 2

(6.18)

The reaction proceeds in two steps during which an atom of copper bound to the en-
zyme is reduced and then oxidized.

E-Cir + -Op * E-Cu© + o 2

(6.19)

E-Cu© + -OP + 2H© -> E-ClT + H 2 0 2 (6.20)

▲ Figure 6.1 1

Surface charge on human superoxide dismu-
tase. The structure of the enzyme is shown
as a model that emphasizes the surface of
the protein. Positively charged regions are
colored blue and negatively charged regions
are colored red. The copper atom at the
active site is green. Note that the channel
leading to the binding site is lined with
positively charged residues. [PDB 1HL5]

The overall reaction includes binding of the anionic substrate
molecules, transfer of electrons and protons, and release of the
uncharged products — all very rapid reactions with this en-
zyme. The k cat /K m value for superoxide dismutase at 25°C is
near 2 x 10 9 M -1 s -1 (Table 6.4). This rate is even faster than
that expected for association of the substrate with the enzyme
based on typical diffusion rates.

How can the rate exceed the rate of diffusion? The expla-
nation was revealed when the structure of the enzyme was ex-
amined. An electric field around the superoxide dismutase
active site enhances the rate of formation of the ES complex
about 30-fold. As shown in Figure 6.11, the active-site copper
atom lies at the bottom of a deep channel in the protein. Hy-
drophilic amino acid residues at the rim of the active-site
pocket guide negatively charged *oP to the positively
charged region surrounding the active site. Electrostatic ef-
fects allow superoxide dismutase to bind and remove super-
oxide (radicals) much faster than expected from random
collisions of enzyme and substrate.

There are probably many enzymes with enhanced rates of
binding due to electrostatic effects. In most cases, the rate-lim-
iting step is catalysis so the overall rate ( k cat /K m ) is slower than
the maximum for a diffusion-controlled reaction. For those
enzymes with fast catalytic reactions, natural selection might favor rapid binding to en-
hance the overall rate. Similarly, an enzyme with rapid binding might evolve a mecha-
nism that favored a faster reaction. However, most biochemical reactions proceed at
rates that are more than sufficient to meet the needs of the cell.

6.5 Modes of Enzymatic Catalysis

The quantitative effects of various catalytic mechanisms are difficult to assess. We have
already seen two chemical mechanisms of enzymatic catalysis, acid-base catalysis and
covalent catalysis. From studies of nonenzymatic catalysts it is estimated that acid-base
catalysis can accelerate a typical enzymatic reaction by a factor of 10 to 100. Covalent
catalysis can provide about the same rate acceleration.

176 CHAPTER 6 Mechanisms of Enzymes

▲ Figure 6.12

Substrate binding. Di hydrofolate reductase
binds NADP + (left) and folate (right), posi-
tioning them in the active site in preparation
for the reductase reaction. Most of the
catalytic rate enhancement is due to binding
effects. [PDB 7DFR]

▲ Figure 6.13

The proximity effect. The enzyme fructose-1, 6-
b/sphosphate aldolase catalyzes the biosyn-
thesis of fructose-1, 6-b/sphosphate from
DHAP and G3P during gluconeogenesis and
the cleavage of fructose-1, 6-b/sphosphate to
dihydroxyacetone phosphate (DHAP) and
glyceraldehyde-3-phosphate (G3P) during
glycolysis (see Section 11.2#4). In the
biosynthesis reaction, the two substrates
DHAP and G3P must be positioned close
together in the active site in an orientation
that promotes their joining to form the larger
fructose-1, 6-b/'sphosphate. This proximity
effect is illustrated for the aldolase from
Mycobacterium tuberculosis. [PDB 2EKZ]

As important as these chemical modes are, they account for only a small portion
of the observed rate accelerations achieved by enzymes (typically 10 8 to 10 12 ). The
ability of proteins to specifically bind and orient ligands explains the remainder.
The proper binding of reactants in the active sites of enzymes provides not only sub-
strate and reaction specificity but also most of the catalytic power of enzymes
(Figure 6.12).

There are two catalytic modes based on binding phenomena. First, for multisub-
strate reactions the collecting and correct positioning of substrate molecules in the
active site raises their effective concentrations over their concentrations in free solution.
In the same way, binding of a substrate near a catalytic active-site residue decreases the
activation energy by reducing the entropy while increasing the effective concentrations
of these two reactants. High effective concentrations favor the more frequent formation
of transition states. This phenomenon is called the proximity effect. Efficient catalysis re-
quires fairly weak binding of reactants to enzymes since extremely tight binding would
inhibit the reaction.

The second major catalytic mode arising from the ligand-enzyme interaction is the
increased binding of transition states to enzymes compared to the binding of substrates
or products. This catalytic mode is called transition state stabilization. There is an equi-
librium (not the reaction equilibrium) between ES and the enzymatic transition state,
ES*. Interaction between the enzyme and its ligands in the transition state shifts this
equilibrium toward ES* and lowers the activation energy.

The effects of proximity and transition- state stabilization were illustrated in Figure 6.3.
Experiments suggest that proximity can increase reaction rates more than 10,000-fold,
and transition-state stabilization can increase reaction rates at least that much. Enzymes
can achieve extraordinary rate accelerations when both of these effects are multiplied by
chemical catalytic effects.

The binding forces responsible for formation of ES complexes and for stabilization
of ES* are familiar from Chapters 2 and 4. These weak forces are charge-charge interac-
tions, hydrogen bonds, hydrophobic interactions, and van der Waals forces. Charge-charge
interactions are stronger in nonpolar environments than in water. Because active sites
are largely nonpolar, charge-charge interactions in the active sites of enzymes can be
quite strong. The side chains of aspartate, glutamate, histidine, lysine, and arginine
residues provide negative and positive groups that form ion pairs with substrates in
active sites. Next in bond strength are hydrogen bonds that often form between
substrates and enzymes. The peptide backbone and the side chains of many amino
acids can form hydrogen bonds. Highly hydrophobic amino acids, as well as alanine,
proline, tryptophan, and tyrosine, can participate in hydrophobic interactions with
the nonpolar groups of ligands. Many weak van der Waals interactions also help bind
substrates. Keep in mind that both the chemical properties of the amino acid residues
and the shape of the active site of an enzyme determine which substrates will bind.

A. The Proximity Effect

Enzymes are frequently described as entropy traps — agents that collect highly mobile
reactants from dilute solution thereby decreasing their entropy and increasing the prob-
ability of their interaction. You can think of the reaction of two molecules positioned at
the active site as an intramolecular (unimolecular) reaction. The correct positioning of
two reacting groups in the active site reduces their degrees of freedom and produces a
large loss of entropy sufficient to account for a large rate acceleration (Figure 6.13). The
acceleration is expressed in terms of the enhanced relative concentration, called the
effective molarity , of the reacting groups in the unimolecular reaction. The effective mo-
larity can be obtained from the ratio

Effective molarity

Ms" 1 )

k 2 ( M _1 s' 1 )

( 6 . 21 )

6.5 Modes of Enzymatic Catalysis 177

where k x is the rate constant when the reactants are preassembled into a single molecule
and k 2 is the rate constant of the corresponding bimolecular reaction. All the units in this
equation cancel except M, so the ratio is expressed in molar units. Effective molarities are
not real concentrations; in fact, for some reactions the values are impossibly high. Never-
theless, effective molarities indicate how favorably reactive groups are oriented.

The importance of the proximity effect is illustrated by experiments comparing
a nonenzymatic bimolecular reaction to a series of chemically similar intramolecular
reactions (Figure 6.14). The bimolecular reaction was the two-step hydrolysis of
p-bromophenyl acetate, catalyzed by acetate and proceeding via the formation of
acetic anhydride. (The second step, hydrolysis of acetic anhydride, is not shown in
Figure 6.14.) In the unimolecular version, reacting groups were connected by a bridge
with progressively greater restriction of rotation. With each restriction placed on
the substrate molecules, the relative rate constant (ki/k 2 ) increased markedly. The glu-
tarate ester (compound 2) has two bonds that allow rotational freedom whereas the
succinate ester (compound 3) has only one. The most restricted compound, the rigid
bicyclic compound 4, has no rotational freedom. In this compound, the carboxylate is

v Figure 6.14

Reactions of a series of carboxylates with
substituted phenyl esters. The proximity
effect is illustrated by the increase in rate
observed when the reactants are held more
rigidly in proximity. Reaction 4 is 50 million
times faster than Reaction 1, the bimolecu-
lar reaction. [Based on Bruice and Pandit
(1960). Intramolecular models depicting
the kinetic importance of “fit” in enzymatic
catalysis. Biochem. 46:402-404.]

Reaction

hUC — Cc~0

Cu^r\

h 3 c — c — o

Relative rate
constants

2 .

H 2 C

H 2 C — C — O u

h 2 c — c
7 \

h 2 c o

\ /

h 2 c — c

1 x 10 3

h 2 c^ c \

h 2 L c / C

2 x 10 5

178 CHAPTER 6 Mechanisms of Enzymes

KEY CONCEPT

The correct binding and positioning of
specific substrates in the active site of an
enzyme produces a large acceleration in
the rate of a reaction.

close to the ester and the reacting groups are properly aligned. The effective molarity of
the carboxylate group is 5 x 10 7 M. Compound 4 has an extremely high probability of
reaction because very little entropy must be lost to reach the transition state. Theoreti-
cal considerations suggest that the greatest rate acceleration that can be expected from
the proximity effect is about 10 8 . This entire rate acceleration can be attributed to the
loss of entropy that occurs when two reactants are properly positioned for reaction.
These intramolecular reactions can serve as a model of the positioning of two substrates
bound in the active site of an enzyme.

B. Weak Binding of Substrates to Enzymes

Reactions of ES complexes are analogous to unimolecular reactions even when two
substrates are involved. Although the correct positioning of substrates in an active site
produces a large rate acceleration, enzymes do not achieve the maximum 10 8 accelera-
tion theoretically generated by the proximity effect. Typically, the loss in entropy on
binding of the substrate allows an acceleration of only 10 4 . That’s because in ES com-
plexes the reactants are brought toward, but not extremely close to, the transition state.
This conclusion is based on both mechanistic reasoning and measurements of the
tightness of binding of substrates and inhibitors to enzymes. One major limitation is
that binding of substrates to enzymes cannot be extremely tight; that is, K m values can-
not be extremely low.

Figure 6.15 shows energy diagrams for a nonenzymatic unimolecular reaction and
the corresponding multistep enzyme -catalyzed reaction. As we will see in the next sec-
tion, an enzyme increases the rate of a reaction by stabilizing (i.e., tightly binding) the
transition state. Therefore, the energy required for ES to reach the transition state (ES*)
in the enzymatic reaction is less than the energy required for S to reach S*, the transition
state in the nonenzymatic reaction.

Recall that the substrate must be bound fairly weakly in the ES complex. If a sub-
strate were bound extremely tightly, it could take just as much energy to reach ES* from
ES (the arrow labeled 2) as is required to reach S* from S in the nonenzymatic reaction
(the arrow labeled 1). In other words, extremely tight binding of the substrate would
mean little or no catalysis. Excessive ES stability is a thermodynamic pit. The role of en-
zymes is to bind and position substrates before the transition state is reached but not so
tightly that the ES complex is too stable.

The K m values (representing dissociation constants) of enzymes for their substrates
show that enzymes avoid the thermodynamic pit. Most K m values are on the order of
10 -4 M, a number that indicates weak binding of the substrate. Enzymes specific for
small substrates, such as urea, carbon dioxide, and superoxide anion, exhibit relatively
high K m values for these compounds (10 -3 to 10 -4 M) because these molecules can
form few noncovalent bonds with enzymes. Enzymes typically have low K m values

Figure 6.15 ►

Energy of substrate binding. In this hypotheti-
cal reaction, the enzyme accelerates the
rate of the reaction by stabilizing the transi-
tion state. In addition, the activation barrier
for formation of the transition state ES*
from ES must be relatively low. If the en-
zyme bound the substrate too tightly
(dashed profile), the activation barrier (2)
would be comparable to the activation bar-
rier of the nonenzymatic reaction (1).

Reaction coordinates

6.5 Modes of Enzymatic Catalysis 179

(10 -6 to 10 -5 M) for coenzymes, which are bulkier than many substrates. The K m values
for the binding of ATP to most ATP- requiring enzymes are about 1(T 4 M or greater but
the muscle-fiber protein myosin (which is not an enzyme) binds ATP a billionfold more
avidly. This large difference in binding reflects the fact that in an ES complex not all
parts of the substrate are bound.

When the concentration of a substrate inside a cell is below the K m value of its corre-
sponding enzyme, the equilibrium of the binding reaction E + S v ES favors E + S. In
other words, the formation of the ES complex is slightly uphill energetically (Figures 6.3
and 6.15), and the ES complex is closer to the energy of the transition state than the
ground state is. This weak binding of substrates accelerates reactions. K m values ap-
pear to be optimized by evolution for effective catalysis — low enough that proximity is
achieved, but high enough that the ES complex is not too stable. The weak binding of
substrates is an important feature of another major force that drives enzymatic catalysis —
increased binding of reactants in the ES^ transition state.

C. Induced Fit

Enzymes resemble solid catalysts by having limited flexibility but they are not entirely
rigid molecules. The atoms of proteins are constantly making small, rapid motions, and
small conformational adjustments occur on binding of ligands. An enzyme is most
effective if it is in the active form initially so no binding energy is consumed in convert-
ing it to an active conformation. In some cases, however, enzymes undergo major shape
alterations when substrate molecules bind. The enzyme shifts from an inactive to an
active form. Activation of an enzyme by a substrate-initiated conformation change is
called induced fit. Induced fit is not a catalytic mode but primarily a substrate specificity
effect.

One example of induced fit is seen with hexokinase, an enzyme that catalyzes the
phosphorylation of glucose by ATP:

Glucose + ATP Glucose 6-phosphate + ADP (6.22)

Water (HOH), which resembles the alcoholic group at C-6 of glucose (ROH), is small
enough and of the proper shape to fit into the active site of hexokinase and therefore it
should be a good substrate. If water entered the active site, hexokinase would quickly
catalyze the hydrolysis of ATP. However, hexokinase -catalyzed hydrolysis of ATP was
shown to be 40,000 times slower than phosphorylation of glucose.

How does the enzyme avoid nonproductive hydrolysis of ATP in the absence of
glucose? Structural experiments with hexokinase show that the enzyme exists in two
conformations: an open form when glucose is absent, and a closed form when glucose is
bound. The angle between the two domains of hexokinase changes considerably when
glucose binds, closing the cleft in the enzyme-glucose complex (Figure 6.16). Produc-
tive hydrolysis of ATP can only take place in the closed form of the enzyme where the
newly formed active site is already occupied by glucose. Water is not a large enough sub-
strate to induce a change in the conformation of hexokinase and this explains why
water does not stimulate ATP hydrolysis. Thus, sugar- induced closure of the hexokinase
active site prevents wasteful hydrolysis of ATP. A number of other kinases follow
induced fit mechanisms.

The substrate specificity that occurs with the induced fit mechanism of hexokinase
economizes cellular ATP but exacts a catalytic price. The binding energy consumed in
moving the protein molecule into the closed shape — a less-favored conformation — is
energy that cannot be used for catalysis. Consequently, an enzyme that uses an induced
fit mechanism is less effective as a catalyst than a hypothetical enzyme that is always in
an active shape and catalyzes the same reaction. The catalytic cost of induced fit slows
kinases so that their /c cat values are approximately 10 3 s -1 (Table 5.1). We will see an-
other example of induced fit and how it economizes metabolic energy in Section 13.3#1
when we describe citrate synthase. The loop-closing reaction of triose phosphate iso-
merase is also an example of an induced fit binding mechanism.

The meaning of K m is discussed in
Section 5.3C. In most cases, it repre-
sents a good approximation of the
dissociation constant for the reaction
E + S ES. Thus, a K m of 10” 4 M
means that at equilibrium the concen-
tration of ES will be approximately
10,000-fold higher than the concentra-
tion of free substrate.

▲ Figure 6.16

Yeast hexokinase. Yeast hexokinase contains
two structural domains connected by a
hinge region. On binding of glucose, these
domains close, shielding the active site from
water, (a) Open conformation, (b) Closed
conformation. [PDB 2YHX and 1HKG].

180 CHAPTER 6 Mechanisms of Enzymes

KEY CONCEPT

Most enzymes exhibit some form of
induced fit binding mechanism.

Hexokinase, citrate synthase, and triose phosphate isomerase are extreme exam-
ples of induced fit mechanisms. Recent advances in the study of enzyme structures re-
veal that almost all enzymes undergo some conformational change when substrate
binds. The simple concept of a rigid lock and a rigid key is being replaced by a more dy-
namic interaction where both the “lock” (enzyme) and the “key” (substrate) adjust to
each other to form a perfect match.

KEY CONCEPT

The catalytic power of enzymes is
explained by binding effects (positioning
the substrates together in the correct
orientation) and stabilization of the
transition state. The result is a lower
activation energy and an increased rate
of reaction.

The role of adenosine deaminase is
described in Section 18.8.

D. Transition-State Stabilization

Enzymes catalyze reactions by physically or electronically distorting the structures of
substrates making them similar to the transition state of the reaction. Transition- state
stabilization — the increased interaction of the enzyme with the substrate in the transi-
tion state — explains a large part of the rate acceleration of enzymes.

Recall Emil Fischer’s lock-and-key theory of enzyme specificity described in Sec-
tion 5.2B. Fischer proposed that enzymes were rigid templates that accepted only cer-
tain substrates as keys. This idea has been replaced by a more dynamic model where
both enzyme and substrate change conformations when they interact. Furthermore,
the classic lock-and-key model dealt with the interaction between enzyme and sub-
strate but we now think of it in terms of enzyme and transition state — the “key” in the
“lock” is the transition state and not the substrate molecule. When a substrate binds to
an enzyme the enzyme distorts the structure of the substrate forcing it toward the
transition state. Maximal interaction with the substrate molecule occurs only in ES^. A
portion of this binding in ES^ can be between the enzyme and nonreacting portions of
the substrate.

An enzyme must be complementary to the transition state in shape and in chemi-
cal character. The graph in Figure 6.15 shows that tight binding of the transition state to
an enzyme can lower the activation energy. Because the energy difference between E + S
and ES^ is significantly less than the energy difference between S and S*, fc cat is greater
than k n (the rate constant for the nonenzymatic reaction). The enzyme-substrate tran-
sition state (ES*) is lower in absolute energy — and therefore more stable — than the tran-
sition state of the reactant in the uncatalyzed reaction. Some transition states may bind
to their enzymes 10 10 to 10 15 times more tightly than their substrates do. The affinity of
other enzymes for their transition states need not be that extreme. A major task for bio-
chemists is to show how transition state stabilization occurs.

The comparative stabilization of ES^ could occur if an enzyme has an active site
with a shape and an electrostatic structure that more closely fits the transition state than
the substrate. An undistorted substrate molecule would not be fully bound. For exam-
ple, an enzyme could have sites that bind the partial charges present only in the unstable
transition state.

Transition- state molecules are ephemeral — they have very short half-lives and are
difficult to detect. One way in which biochemists can study transition states is to create
stable analogs that can bind to the enzyme. These transition-state analogs are molecules
whose structures resemble presumed transition states. If enzymes prefer to bind to tran-
sition states, then a transition- state analog should bind extremely tightly to the appro-
priate enzyme — much more tightly than substrate — and thus be a potent inhibitor. The
dissociation constant for a transition state analog should be about 10 -13 M or less.

One of the first examples of a transition-state analog was 2-phosphoglycolate
(Figure 6.17), whose structure resembles the first enediolate transition state in the reac-
tion catalyzed by triose phosphate isomerase (Section 6.4A). This transition- state ana-
log binds to the isomerase at least 100 times more tightly than either of the substrates of
the enzyme (Figure 6.18). Tighter binding results from a partially negative oxygen atom
in the carboxylate group of 2-phosphoglycolate, a feature shared with the transition
state but not with the substrates.

Experiments with adenosine deaminase have identified a transition- state analog
that binds to the enzyme with amazing affinity because it resembles the transition state
very closely. Adenosine deaminase catalyzes the hydrolytic conversion of the purine nu-
cleoside adenosine to inosine. The first step of this reaction is the addition of a molecule

6.5 Modes of Enzymatic Catalysis 181

H0 \ /Opo 3 ©

C CH2

H H

H0 \^ c \ /OPo 3 ©
C CH2

Dihydroxyacetone

phosphate

Transition state

opo 3 ©

O CH2

2-Phosphoglycolate
(transition-state analog)

I ^

HO^ ^0P0 3 ©

CH2

Enediol intermediate

◄ Figure 6.17

2-Phosphoglycolate, a transition-state
analog for the enzyme triose phosphate
isomerase. 2-Phosphoglycolate is pre-
sumed to be an analog of C-2 and C-3
of the transition state (center) between
dihydroxyacetone phosphate (right) and
the initial enediolate intermediate in
the reaction.

of water (Figure 6.19a). The complex with water, called a covalent hydrate, forms as soon
as adenosine is bound to the enzyme and quickly decomposes to products. Adenosine
deaminase has broad substrate specificity and catalyzes the hydrolytic removal of vari-
ous groups from position 6 of purine nucleosides. However, the inhibitor purine ri-
bonucleoside (Figure 6.19b) has just hydrogen at position 6 and undergoes only the first
enzymatic step of hydrolysis, addition of the water molecule. The covalent hydrate that’s
formed is a transition- state analog, a competitive inhibitor having a FQ of 3 X 10 -13 M.
(For comparison, the affinity constant of adenosine deaminase for its true transition
state is expected to be 3 X 10 -17 M.). The binding of this analog exceeds the binding of
either the substrate or the product by a factor of more than 10 8 . A very similar reduced
inhibitor, 1,6-dihydropurine ribonucleoside (Figure 6.19c), lacks the hydroxyl group at
C-6, and it has a K x of only 5 x 10~ 6 M. We can conclude from these studies that adenosine

◄ Figure 6.18

Binding of 2-phosphoglycolate to triose phos-
phate isomerase. The transition state ana-
logue, 2-phosphoglycolate is bound at the
active site of Plasmodium falciparum triose
phosphate isomerase. The molecule is held
in position by many hydrogen bonds between
the phosphate group and surrounding amino
acid side chains. Some of the hydrogen
bonds are formed through bridged “frozen”
water molecules in the active site. The
catalytic residues, Glu-165 and His-95,
form hydrogen bonds with the carboxylate
group of 2-phosphoglycolate as expected in
the transition state. [PDB 1LYZ]

Glu 165

182 CHAPTER 6 Mechanisms of Enzymes

H 2 N OH

Ribose

Adenosine Covalent hydrate

(substrate)

Inosine

(product)

(c)

Ribose

Purine ribonucleoside
(substrate analog)

Transition-state

analog

1,6-Dihydropurine ribonucleoside
(competitive inhibitor)

▲ Figure 6.20

Adenosine deaminase with bound transition-
state analog.

▲ Figure 6.19

Inhibition of adenosine deaminase by a transition-state analog, (a) In the deamination
of adenosine, a proton is added to N-l and a hydroxide ion is added to C-6 to form
an unstable covalent hydrate, which decomposes to produce inosine and ammonia,
(b) The inhibitor purine ribonucleoside also rapidly forms a covalent hydrate, 6-hy-
droxy-l,6-dihydropurine ribonucleoside. This covalent hydrate is a transition-state
analog that binds more than a million times more avidly than another competitive
inhibitor, 1,6-dihydropurine ribonucleoside (c), which differs from the transition-
state analog only by the absence of the 6-hydroxyl group.

deaminase must specifically and avidly bind the transition-state analog —
and also the transition state — through interaction with the hydroxyl
group at C-6.

The structure of adenosine deaminase with the bound transition-
state analog is shown in Figure 6.20 and the interactions between the
analog and amino acid side chains in the active site are depicted in Figure
6.21. Notice the hydrogen bonds between Asp-292 and the hydroxyl
group on C-6 of 6-hydroxy- 1,6-dihydropurine and the interaction be-
tween this hydroxyl group and a bound zinc ion in the active site. This
confirms the hypothesis that the enzyme specifically binds the transition
state in the normal reaction.

His14

Asp16

His12 Hisl 5

Asp296 \ / H j s2 1 1

^ Zn 2 T.

N HO— -

\ NH-__

N=/

-OH

Asp292

-Glu214

.OH OH

\ / NH^

Wat 569 G ^ 81 ^

▲ Figure 6.21

Transition-state analog binding to adenosime deaminase. The interactions between the transition state
analog, 6-hydroxy-l,6-dihydropurine, and amino acid side chains in the active site of adenosine
deaminase confirms that the enyme recognizes the hydroxyl group at C-6. [PDB 1KRM]

6.6 Serine Proteases

183

6.6 Serine Proteases

Serine proteases are a class of enzymes that cleave the peptide bond of proteins. As the
name implies, they are characterized by the presence of a catalytic serine residue in their
active sites. The best-studied serine proteases are the related enzymes trypsin, chy-
motrypsin, and elastase. These enzymes provide an excellent opportunity to explore the
relationship between protein structure and catalytic function. They have been inten-
sively studied for 50 years and form an important part of the history of biochemistry
and the elucidation of enzyme mechanisms. In this section, we see how the activity of
serine proteases is regulated by zymogen activation and examine a structural basis for
the substrate specificity of different serine proteases.

A. Zymogens Are Inactive Enzyme Precursors

Mammals digest food in the stomach and intestines. During this process, food pro-
teins undergo a series of hydrolytic reactions as they pass through the digestive tract.
Following mechanical disruption by chewing and moistening with saliva, foods are
swallowed and mixed with hydrochloric acid in the stomach. The acid denatures pro-
teins and pepsin (a protease that functions optimally in an acidic environment) cat-
alyzes hydrolysis of these denatured proteins to a mixture of peptides. The mixture
passes into the intestine where it is neutralized by sodium bicarbonate and digested
by the action of several proteases to amino acids and small peptides that can be ab-
sorbed into the bloodstream.

Pepsin is initially secreted as an inactive precursor called pepsinogen. When
pepsinogen encounters HC1 in the stomach it is activated to cleave itself forming the
more active protease, pepsin. The stomach secretions are stimulated by food — or even
the anticipation of food — as shown by Ivan Pavlov in his experiments with dogs over
100 years ago. (Pavlov was awarded a Nobel Prize in 1904.) The inactive precursor is
called a zymogen. Pavlov was the first to show that zymogens could be converted to ac-
tive proteases in the stomach and intestines.

The main serine proteases are trypsin, chymotrypsin, and elastase. Together, they
catalyze much of the digestion of proteins in the intestine. Like pepsin, these enzymes are
also synthesized and stored as inactive precursors called zymogens. The zymogens, are called
trypsinogen, chymotrypsinogen, and proelastase. They are synthesized in the pancreas.
Its important to store these hydrolytic enzymes as inactive precursors within the cell since
the active proteases would kill the pancreatic cells by cleaving cytoplasmic proteins.

BOX 6.3 KORNBERG’S TEN COMMANDMENTS

1. Rely on enzymology to clarify biologic questions

2. Trust the universality of biochemistry and the power of microbiology

3. Do not believe something because you can explain it

4. Do not waste clean thinking on dirty enzymes

5. Do not waste clean enzymes on dirty substrates

6. Depend on viruses to open windows

7. Correct for extract dilution with molecular crowding

8. Respect the personality of DNA

9. Use reverse genetics and genomics

10. Employ enzymes as unique reagents

Arthur Kornberg, Nobel Laureate in Physiology or Medicine 1959

Kornberg, A. (2000). Ten commandments: lessons from the enzymology of DNA replication.
/. Bacteriol. 182:3613-3618.

Kornberg, A. (2003). Ten commandments of enzymology, amended. Trends Biochem. Sci.
28:515-517.

184 CHAPTER 6 Mechanisms of Enzymes

Trypsinogen

Enteropeptidase

Trypsin ^

Chymotrypsinogen

◄

V +

Chymotrypsin

i Proelastase

◄ x v — ►

+ +

Elastase

▲ Figure 6.22

Activation of some pancreatic zymogens.

Initially, enteropeptidase catalyzes the acti-
vation of trypsinogen to trypsin. Trypsin then
activates chymotrypsinogen, proelastase,
and additional trypsinogen molecules.

(a)

(b)

▲ Figure 6.23

Polypeptide chains of chymotrypsinogen (left)
[PDB 2CGA] and a-chymotrypsin (right) [PDB
5CHA]. lle-16 and Asp-194 in both zymogen
and the active enzyme are shown in yellow.
The catalytic-site residues (Asp-102, His-
57, and Ser-195) are shown in red. The
residues that are removed by processing the
zymogen are colored green.

The enzymes are activated by selective proteolysis — enzymatic cleavage of one or a
few specific peptide bonds — when they are secreted from the pancreas into the small in-
testine. A protease called enteropeptidase specifically activates trypsinogen to trypsin by
catalyzing cleavage of the bond between Lys-6 and Ile-7. Once activated by the removal of
its N-terminal hexapeptide, trypsin proteolytically cleaves the other pancreatic zymogens,
including additional trypsinogen molecules (Figure 6.22).

The activation of chymotrypsinogen to chymotrypsin is catalyzed by trypsin and
by chymotrypsin itself. Four peptide bonds (between residues 13 and 14, 15 and 16, 146
and 147, and 148 and 149) are cleaved resulting in the release of two dipeptides. The result-
ing chymotrypsin retains its three-dimensional shape, despite two breaks in its back-
bone. This stability is partly due to the presence of five disulfide bonds in the protein.

X-ray crystallography has revealed one major difference between the conformation
of chymotrypsinogen and chymotrypsin — the lack of a hydrophobic substrate-binding
pocket in the zymogen. The differences are shown in Figure 6.23 where the structures of
chymotrypsinogen and chymotrypsin are compared. On zymogen activation, the newly
generated a-amino group of lie- 16 turns inward and interacts with the /3 -carboxyl
group of Asp- 194 to form an ion pair. This local conformational change generates a rel-
atively hydrophobic substrate-binding pocket near the three catalytic residues with ion-
izable side chains (Asp- 102, His- 57, and Ser-195).

B. Substrate Specificity of Serine Proteases

Chymotrypsin, trypsin, and elastase are similar enzymes that share a common ancestor;
in other words, they are homologous. Each enzyme has a two-lobed structure with the
active site located in a cleft between the two domains. The positions of the catalytically
active side chains of the serine, histidine, and aspartate residues in the active sites are
almost identical in the three enzymes (Figure 6.24).

The substrate specificities of chymotrypsin, trypsin, and elastase have been
explained by relatively small structural differences in the enzymes. Recall that trypsin
catalyzes the hydrolysis of peptide bonds whose carbonyl groups are contributed by
arginine or lysine (Section 3.10). Both chymotrypsin and trypsin contain a binding
pocket that correctly positions the substrates for nucleophilic attack by an active-site
serine residue. Each protease has a similar extended region into which polypeptides fit
but the so-called specificity pocket near the active- site serine is markedly different for
each enzyme. Trypsin differs from chymotrypsin because in chymotrypsin there is an
uncharged serine residue at the base of the hydrophobic binding pocket. In trypsin this
residue is an aspartate residue (Figure 6.25). This negatively charged aspartate residue
forms an ion pair with the positively charged side chain of an arginine or lysine residue
of the substrate in the ES complex. Experiments with specifically mutated trypsin indicate
that the aspartate residue at the base of its specificity pocket is a major factor in sub-
strate specificity but other parts of the molecule also affect specificity.

Elastase catalyzes the degradation of elastin, a fibrous protein that is rich in glycine
and alanine residues. Elastase is similar in tertiary structure to chymotrypsin except that

(a) (b) (c)

▲ Figure 6.24

Serine proteases. Comparison of the polypeptide backbones of (a) chymotrypsin [PDB 5CHA], (b) trypsin
[PDB 1TLD], and (c) elastase [PDB 3EST]. Residues at the catalytic center are shown in red.

6.6 Serine Proteases 185

(a) Chymotrypsin

Ser

(b) Trypsin

• Carbon
O Nitrogen

# Oxygen

◄ Figure 6.25

Binding sites of chymotrypsin, trypsin, and
elastase. The differing binding sites of these
three serine proteases are primary determinants
of their substrate specificities, (a) Chymotrypsin
has a hydrophobic pocket that binds the side
chains of aromatic or bulky hydrophobic amino
acid residues, (b) A negatively charged as-
partate residue at the bottom of the binding
pocket of trypsin allows trypsin to bind the
positively charged side chains of lysine and
arginine residues, (c) In elastase, the side
chains of a valine and a threonine residue at
the binding site create a shallow binding
pocket. Elastase binds only amino acid
residues with small side chains, especially
glycine and alanine residues.

its binding pocket is much shallower. Two glycine residues found at the entrance of the
binding site of chymotrypsin and trypsin are replaced in elastase by much larger valine
and threonine residues (Figure 6.25c). These residues keep potential substrates with
large side chains away from the catalytic center. Thus, elastase specifically cleaves pro-
teins that have small residues such as glycine and alanine.

C. Serine Proteases Use Both the Chemical
and the Binding Modes of Catalysis

Let s examine the mechanism of chymotrypsin and the roles of three catalytic residues:
His-57, Asp- 102, and Ser- 195. Many enzymes catalyze the cleavage of amide or ester
bonds by the same process so study of the chymotrypsin mechanism can be applied to a
large family of hydrolases.

Asp- 102 is buried in a rather hydrophobic environment. It is hydrogen-bonded to
His-57 that in turn is hydrogen-bonded to Ser- 195 (Figure 6.26 ). This group of amino acid
residues is called the catalytic triad. The reaction cycle begins when His-57 abstracts a pro-
ton from Ser-195 (Figure 6.27). This creates a powerful nucleophile (Ser-195) that will
eventually attack the peptide bond. Initiation of this part of the reaction is favored because
Asp- 102 stabilizes the histidine promoting its ability to deprotonate the serine residue.

The discovery that Ser-195 is a catalytic residue of chymotrypsin was surprising be-
cause the side chain of serine is usually not sufficiently acidic to undergo deprotonation
in order to serve as a strong nucleophile. The hydroxymethyl group of a serine residue
generally has a p K a of about 16 and is similar in reactivity to the hydroxyl group of
ethanol. You may recall from organic chemistry that although ethanol can ionize to

▲ Figure 6.26

The catalytic site of chymotrypsin. The active-
site residues Asp-102, His-57, and Ser-195
are arrayed in a hydrogen-bonded network.
The conformation of these three residues is
stabilized by a hydrogen bond between the
carbonyl oxygen of the carboxylate side
chain of Asp-102 and the peptide-bond
nitrogen of His-57. Oxygen atoms of the
active-site residues are red, and nitrogen
atoms are dark blue. [PDB 5CHA].

His-57

Asp-

102

ch 2

Asp-

102

CH,

-ch 2

, 0 +

.H^ ^ ^H|

Ser-195

ch 2

▲ Figure 6.27

Catalytic triad of chymotrypsin. The imidazole ring of His-57 removes the proton from the hydroxymethyl side chain of Ser-195 (to
which it is hydrogen-bonded), thereby making Ser-195 a powerful nucleophile. This interaction is facilitated by interaction of the
imidazolium ion with its other hydrogen-bonded partner, the buried /3-carboxylate group of Asp-102. The residues of the triad are
drawn in an arrangement similar to that shown in Figure 6.24.

186 CHAPTER 6 Mechanisms of Enzymes

BOX 6.4 CLEAN CLOTHES

Its a little-known fact that 75% of all laundry detergents contain
proteases that are used in helping to remove stubborn protein-
based stains from dirty clothes.

All protease additives are based on serine proteases iso-
lated from various Bacillus species. These enzymes have been
extensively modified in order to be active under the harsh
conditions of a detergent solution at high temperature. A
successful example of site-directed mutagenesis is the alter-
ation of the serine protease subtilisin from Bacillus subtilis
(Box 6.4) to make it more resistant to chemical oxidation. It
has a methionine residue in the active-site cleft (Met-222)
that is readily oxidized leading to inactivation of the enzyme.
Resistance to oxidation increases the suitability of subtilisin
as a detergent additive. Met-222 was systematically replaced
by each of the other common amino acids in a series of mu-
tagenic experiments. All 19 possible mutant subtilisins were
isolated and tested and most had greatly diminished peptidase
activity. The Cys-222 mutant had high activity but was also
subject to oxidation. The Ala-222 and Ser-222 mutants, with
nonoxidizable side chains, were not inactivated by oxidation
and had relatively high activity. They were the only active,
oxygen- stable mutant subtilisin variants.

Site-directed mutagenesis has been performed to alter
eight of the 319 amino acid residues of a bacterial protease.

The wild-type protease is moderately stable when heated but
the suitably mutated enzyme is stable and can function at
100°C. Its denaturation in detergent is prevented by groups,
such as a disulfide bridge, that stabilize its conformation.

Recently there has been a trend to lower wash tempera-
tures in order to save energy. The older group of enzymes are
not effective at lower wash temperatures so a whole new
round of bioengineering has begun creating modified en-
zymes that can be effective in a modern energy-conscious
household.

form an ethoxide this reaction requires the presence of an extremely strong base or
treatment with an alkali metal. We see below how the active site of chymotrypsin,
achieves this ionization in the presence of a substrate.

A proposed mechanism for chymotrypsin and related serine proteases includes co-
valent catalysis (by a nucleophilic oxygen) and general acid-base catalysis (donation of
a proton to form a leaving group). The steps of the proposed mechanism are illustrated
in Figure 6.28.

Binding of the peptide substrate causes a slight conformation change in chy-
motrypsin, sterically compressing Asp- 102 and His-57. A low-barrier hydrogen bond is
formed between these side chains and the p K a of His-57 rises from about 7 to about 11.
(Formation of this strong, almost covalent, bond drives electrons toward the second N
atom of the imidazole ring of His-57 making it more basic.) This increase in basicity
makes His-57 an effective general base for abstracting a proton from the — CH 2 OH of
Ser-195. This mechanism explains how the normally unreactive alcohol group of serine
becomes a potent nucleophile.

All the catalytic modes described in this chapter are used in the mechanisms of ser-
ine proteases. In the reaction scheme shown in Figure 6.28, steps 1 and 4 in the forward
direction use the proximity effect, the gathering of reactants. For example, when a water
molecule replaces the amine (Pi) in step 4, it is held by histidine, providing a proximity
effect. Acid-base catalysis by histidine lowers the energy barriers for steps 2 and 4. Co-
valent catalysis using the — CH 2 OH of serine occurs in steps 2 through 5. The unstable
tetrahedral intermediates at steps 2 and 4 (E-TIi and E-TI 2 ) are believed to be similar to
the transition states for these steps. Hydrogen bonds in the oxyanion hole stabilize these
intermediates, which are oxyanion forms of the substrate, by binding them more tightly
to the enzyme than the substrate was bound. The chemical modes of catalysis
(acid-base and covalent catalysis) and the binding modes of catalysis (the proximity ef-
fect and transition-state stabilization) all contribute to the enzymatic activity of serine
proteases.

6.6 Serine Proteases 187

BOX 6.5 CONVERGENT EVOLUTION

The protease subtilisin from the bacterium Bacillus subtilis is
another example of a serine protease. It possesses a catalytic
triad consisting of Asp-32, His-64, and Ser-221 at its active
site. These are arranged in an alignment similar to the Asp- 102,
His-57, and Ser-195 residues in chymotrypsin (Figure 6.27).
However, as you might deduce from the residue numbers,
the structures of subtilisin and chymotrypsin are very differ-
ent and there is no significant sequence similarity.

This is a remarkable example of convergent evolution.
The mammalian intestinal serine proteases and the bacterial
subtilisins have independently discovered the catalytic Asp-
His-Ser triad.

► Subtilisin from Bacillus
subtilis. The structure
of this enzyme is very
different from that of
serine proteases shown in
Figure 6.24. [PDB 1SBC]

6.7 Lysozyme

Lysozyme catalyzes the hydrolysis of some polysaccharides, especially those that make
up the cell walls of bacteria. It is the first enzyme whose structure was solved and for
this reason there has been a long-term interest in working out its precise mechanism of
action. Many secretions, such as tears, saliva, and nasal mucus, contain lysozyme activ-
ity to help prevent bacterial infection. (Lysozyme causes lysis , or disruption, of bacterial
cells.) The best-studied lysozyme is from chicken egg white.

The substrate of lysozyme is a polysaccharide composed of alternating residues
of N-acetylghicosamine (GlcNAc) and N-acetylmuramic acid (MurNAc) connected y^g structure of bacterial cell walls is

by glycosidic bonds (Figure 6.29). Lysozyme specifically catalyzes hydrolysis of the described in Seciton 8 7B

glycosidic bond between C-l of a MurNAc residue and the oxygen atom at C-4 of a
GlcNAc residue.

Models of lysozyme and its complexes with saccharides have been obtained by
X-ray crystallographic analysis (Figure 6.30). The substrate-binding cleft of lysozyme
accommodates six saccharide residues. Each of the residues binds to a particular part of
the active cleft at sites A through E.

Sugar molecules fit easily into all but one site of the structural model. At site D a
sugar molecule such as MurNAc does not fit into the model unless it is distorted into a

Lysozyme

◄ Figure 6.29

Structure of a four-residue portion of a bacter-
ial cell-wall polysaccharide. Lysozyme cat-
alyzes hydrolytic cleavage of the glycosidic
bond between C-l of MurNAc and the oxy-
gen atom involved in the glycosidic bond.

cn 3

c = o

CH 2 OH

GlcNAc

MurNAc

GlcNAc

MurNAc

188 CHAPTER 6 Mechanisms of Enzymes

The noncovalent enzyme-substrate
complex is formed, orienting the
substrate for reaction. Interactions
holding the substrate in place
include binding of the R ^ group in
the specificity pocket (shaded). The
binding interactions position the
carbonyl carbon of the scissile
peptide bond (the bond
susceptible to cleavage) next to
the oxygen of Ser-195.

Binding of the substrate compresses
Asp-102 and His-57. This strain is
relieved by formation of a low-barrier
hydrogen bond. The raised pK a of
His-57 enables the imidazole ring to
remove a proton from the hydroxyl
group of Ser-195. The nucleophilic
oxygen of Ser-195 attacks the
carbonyl carbon of the peptide bond
to form a tetrahedral intermediate
(E-Th), which is believed to resemble
the transition state.

When the tetrahedral intermediate is
formed, the substrate C — O bond
changes from a double bond to a longer
single bond. This allows the negatively
charged oxygen (the oxyanion) of the
tetrahedral intermediate to move to a E-T^
previously vacant position, called the
oxyanion hole, where it can form
hydrogen bonds with the peptide-chain
— NH groups of Gly-193 and Ser-195.

The imidazolium ring of His-57 acts as
an acid catalyst, donating a proton to
the nitrogen of the scissile peptide
bond, thus facilitating its cleavage.

The carbonyl group from the peptide
forms a covalent bond with the
enzyme, producing an acyl-enzyme Acyl E

intermediate. After the peptide +

product (Pt) with the new amino
terminus leaves the active site, water
enters.

Ser-195

▲ Figure 6.28

Mechanism of chymotrypsin-catalyzed cleavage of a peptide bond.

6.7 Lysozyme

189

Ser-195

Carboxylate product (P 2 )

The carboxylate product is released from
the active site, and free chymotrypsin is
regenerated.

The second product (P 2 ) — a polypeptide
with a new carboxy terminus — is formed.

His-57, once again an imidazolium ion,
donates a proton, leading to the collapse
of the second tetrahedral intermediate.

A second tetrahedral intermediate (E-TI 2 )
is formed and stabilized by the oxyanion
hole.

Hydrolysis (deacylation) of the acyl-
enzyme intermediate starts when
Asp-102 and His-57 again form a low-
barrier hydrogen bond and His-57
removes a proton from the water
molecule to provide an OH^group to
attack the carbonyl group of the ester.

▲ Figure 6.28 ( continued )

190 CHAPTER 6 Mechanisms of Enzymes

▲ Figure 6.30

Lysozyme from chicken with a pentasaccharide
molecule (pink). The ligand is bound in sites
A, B, C, D and E. Site F is not occupied
in this structure. The active site for bond
cleavage is between sites D and E.

[PDB 1SFB].

(a) Chair conformation

H | H

0=C

ch 3

(b) Half-chair conformation

o=c

ch 3

▲ Figure 6.31

Conformations of /V-acetylmuramic acid.

(a) Chair conformation, (b) Half-chair con-
formation proposed for the sugar bound in
site D of lysozyme. R represents the lactyl
group of MurNAc.

half-chair conformation (Figure 6.31). Two ionic amino acid residues, Glu-35 and
Asp-52, are located close to C-l of the distorted sugar molecule in the D binding site.
Glu-35 is in a nonpolar region of the cleft and has a perturbed piC a near 6.5. Asp-52, in
a more polar environment, has a piC a near 3.5. The pH optimum of lysozyme is near
5 — between these two p K a values. Recall that the piC a value of individual amino acid
side chains may not be the same as the piC a value of the free amino acid in solution
(Section 3.4).

The proposed mechanism of lysozyme is shown in Figure 6.32. When a molecule of
polysaccharide binds to lysozyme, MurNAc residues bind to sites B, D, and F (there is
no cavity for the lactyl side chain of MurNAc in site A, C, or E). The extensive binding of
the oligosaccharide forces the MurNAc residue in the D site into the half- chair confor-
mation. A near covalent bond forms between Asp-52 and the postulated intermediate
(an unstable oxocarbocation). Recent evidence suggests that this interaction might be
more like a covalent bond than a strong ion pair but there is much controversy over this
point. Its interesting that there are still details of the lysozyme mechanism to be worked
out after almost 50 years of effort.

Lysozyme is only one representative of a large group of glycoside hydrolases. Re-
cently, the structures of a bacterial cellulase and its complexes with substrate, intermediate,
and product have been determined. This glycosidase has a slightly different mecha-
nism than lysozyme — it forms a covalent glycosyl-enzyme intermediate rather than
the strong ion pair postulated for lysozyme. Other aspects of its mechanism, such
as distortion of a sugar residue and interaction with active-site — COOH and
— COO^ side chains, resemble those of the lysozyme mechanism. The structures
of the enzyme complexes show that distortion of the substrate forces it toward the
transition state.

6.8 Arginine Kinase

Most enzymatic reactions for which detailed mechanisms have been elucidated involve
fairly simple reactions, such as isomerizations, cleavage reactions, or reactions with
water as the second reactant. Therefore, in order to assess proximity effects and the ex-
tent of transition state stabilization, it’s worthwhile looking at a more complicated reac-
tion, such as that catalyzed by arginine kinase:

Arginine + MgATP Arginine Phosphate + MgADP + H®

The structure of a transition- state analog-enzyme complex of arginine kinase has
been determined at high resolution (Figure 6.33). However, rather than studying the
usual type of transition-state analog in which reactants are fused by covalent bonds, the
scientists used three separate components: arginine, nitrate (to model the phosphoryl
group transferred between arginine and ADP), and ADP. X-ray crystallographic exami-
nation of the active site containing these three compounds led to the proposal of a
structure for the transition state and a mechanism for the reaction (see Figure 6.33).
The crystallographic results showed that the enzyme has greatly restricted the move-
ment of the bound species (and presumably also of the transition state). For example,
the terminal pyrophosphoryl group of ATP is held in place by four arginine side chains
and a bound Mg 2+ ion and the guanidinium group of the arginine substrate molecule is
held firmly by two glutamate side chains. The components are precisely and properly
aligned by the enzyme.

Arginine kinase, like other kinases, is an induced-fit enzyme (Section 6.5C). It as-
sumes the closed shape when it is crystallized in the presence of arginine, nitrate, and
ADP. This enzyme has a k cat of about 2 x 10 2 s -1 and K m values above 10 -4 M for both
arginine and ATP — values that are quite typical for kinases. The movement that occurs
during the induced-fit binding of substrates has precisely aligned the substrates, which
had previously been bound fairly weakly, as shown by their moderate K m values. At least
four interrelated catalytic effects participate in this enzymatic reaction: proximity

6.8 Arginine Kinase 191

A MurNAc residue of the
substrate is distorted when
it binds to the D site.

Glu-35, which is protonated at pH 5,
acts as an acid catalyst, donating a proton
to the oxygen involved in the glycosidic
bond between the the D and E residues.

The portion of the substrate bound

Asp-52, which is negatively
charged at pH 5, forms a strong
ion pair with the unstable
oxocarbocation intermediate.
This interaction is close to a
covalent bond.

A proton from the water molecule is

▲ Figure 6.32

Mechanism of lysozyme. Ri represents the lactyl group, and R 2 represents the A/-acetyl group of MurNAc.

192 CHAPTER 6 Mechanisms of Enzymes

Figure 6.33 ►

Proposed structure of the active site of arginine
kinase in the presence of ATP and arginine.

The substrate molecules are held firmly and
aligned toward the transition state, as shown
by the dashed lines. The asterisks (*) show
that either Glu-225 or Glu-314 could act as
a general acid-base catalyst.

{Adapted from Zhov, G., Somasundaram, T., Blanc,
E., Parthasarathy, G., Ellington, W. R., and Chapman,
M. S. (1998). Transition state structure of arginine
kinase: implications for catalysis of bimolecular
reactions. Proc. Natl. Acad. Sci. USA. 95:8453.)

\ Thr-273 <-/Cys-271

O — H -"' 1 2 3 4 S 6 0

Glu-225.

0-— .
0

/ N

. N -

H-

arginine

\©

Arg-229N® .

Arg-126 7 N — H

H''\-

N u

H 2 N H /

— N

Arg-280\\@
H->N

0^"

: O

N— H-

-O

-:0

Glu-314

H "" 0*

.© H

;;Mg©

>^©

/ °— -

ATP

'NH

(collection and alignment of substrate molecules), fairly weak initial binding of sub-
strates, acid-base catalysis, and transition-state stabilization (strain of substrates toward
the shape of the transition state).

Having gained insight into the general mechanisms of enzymes, we can now go on
to examine reactions that include coenzymes. These reactions require groups not sup-
plied by the side chains of amino acids.

Summary

1. The four major modes of enzymatic catalysis are acid-base catalysis
and covalent catalysis (chemical modes) and proximity and tran-
sition-state stabilization (binding modes). The atomic details of
reactions are described by reaction mechanisms, which are based
on the analysis of kinetic experiments and protein structures.

2. For each step in a reaction, the reactants pass through a transition
state. The energy difference between stable reactants and the tran-
sition state is the activation energy. Catalysts allow faster reactions
by lowering the activation energy.

3. Ionizable amino acid residues in active sites form catalytic cen-
ters. These residues may participate in acid-base catalysis (proton
addition or removal) or covalent catalysis (covalent attachment of
a portion of the substrate to the enzyme). The effects of pH on
the rate of an enzymatic reaction can suggest which residues par-
ticipate in catalysis.

4. The catalytic rates for a few enzymes are so high that they ap-
proach the upper physical limit of reactions in solution, the rate
at which reactants approach each other by diffusion.

5. Most of the rate acceleration achieved by an enzyme arises from
the binding of substrates to the enzyme.

6. The proximity effect is acceleration of the reaction rate due to the
formation of a noncovalent ES complex that collects and orients
reactants resulting in a decrease in entropy.

7. An enzyme binds its substrates fairly weakly. Excessively strong
binding would stabilize the ES complex and slow the reaction.

8. An enzyme binds a transition state with greater affinity than it
binds substrates. Evidence for transition state stabilization is pro-
vided by transition-state analogs that are enzyme inhibitors.

9. Some enzymes use induced fit (substrate-induced activation that
involves a conformation change) to prevent wasteful hydrolysis of
a reactive substrate.

10. Many serine proteases are synthesized as inactive zymogens that
are activated extracellularly under appropriate conditions by se-
lective proteolysis. The examination of serine proteases by X-ray
crystallography shows how the three-dimensional structures of
proteins can reveal information about the active sites, including
the binding of specific substrates.

11. The active sites of serine proteases contain a hydrogen-bonded
Ser-His-Asp catalytic triad. The serine residue serves as a cova-
lent catalyst, and the histidine residue serves as an acid-base cata-
lyst. Anionic tetrahedral intermediates are stabilized by hydrogen
bonds with the enzyme.

12. The proposed mechanism for lysozyme, an enzyme that catalyzes
the hydrolysis of bacterial cell walls, includes substrate distortion
and stabilization of an unstable oxocarbocation intermediate.

Problems 193

Problems

1. (a) What forces are involved in binding substrates and interme-

diates to the active sites of enzymes?

(b) Explain why very tight binding of a substrate to an enzyme is
not desirable for enzyme catalysis, whereas tight binding of
the transition state is desirable.

2. The enzyme orotodine 5-phosphate decarboxylase is one of the
most proficient enzymes known, accelerating the rate of decarboxy-
lation of orotidine 5' monophosphate by a factor of 10 23 (Section
5.4). Nitrogen- 15 isotope effect studies have shown that two major
participating mechanisms are (1) destabilization of the ground state
ES complex by electrostatic repulsion between the enzyme and sub-
strate, and (2) stabilization of the transition state by favorable elec-
trostatic interactions between the enzyme and ES*. Draw an energy
diagram that shows how these two effects promote catalysis.

3. The energy diagrams for two multistep reactions are shown below.
What is the rate- determining step in each of these reactions?

4. Reaction 2 below occurs 2.5 X 10 11 times faster than Reaction 1.
What is likely to be a major reason for this enormous rate increase
in Reaction 2? How is this model relevant for interpreting possi-
ble mechanisms for enzyme rate increases?

5. List three major catalytic effects for lysozyme and explain how
each is used during the enzyme- catalyzed hydrolysis of a glyco-
sidic bond.

6. There are multiple serine residues in a- chymotrypsin but only ser-
ine 195 reacts rapidly when the enzyme is treated with active phos-
phate inhibitors such as diisopropyl fluorophosphate (DFP). Explain.

7. (a) Identify the residues in the catalytic triad of a-chymotrypsin

and indicate the type of catalysis mediated by each residue.

(b) What additional amino acid groups are found in the oxy an-
ion hole and what role do they play in catalysis?

(c) Explain why site-directed mutagenesis of aspartate to as-
paragine in the active site of trypsin decreases the catalytic
activity 10,000-fold.

8. Catalytic triad groupings of amino acid residues increase the nu-
cleophilic character of active-site serine, threonine, or cysteine
residues present in many enzymes involved in catalyzing the cleav-
age of substrate amide or ester bonds. Using a- chymotrypsin as a
model system, diagram the expected arrangements of the catalytic
triads in the enzymes below.

(a) Human cytomegalovirus protease: His, His, Ser

(b) /3- lactamase: Glu, Lys, Ser

(d) Hepatitis A protease: Asp, (H 2 0), His, Cys (a water molecule
is situated between the Asp and His residues)

9 . Human dipeptidyl peptidase IV (DDP-IV) is a serine protease
that catalyzes hydrolysis of prolyl peptide bonds at the next-
to-last position at the N terminus of a protein. Many physiologi-
cal peptides have been identified as substrates, including proteins
involved in the regulation of glucose metabolism. DDP-IV con-
tains a catalytic triad at the active site (Glu-His-Ser) and a tyrosine
residue in the oxyanion hole. Site-directed mutagenesis of this
tyrosine residue in DPP-IV was performed, and the ability of
the enzyme to cleave a peptide substrate was compared to that of the
wild-type enzyme. The tyrosine residue found in the oxyanion
hole was changed to a phenylalanine. The phenylalanine mutant
had less than 1% of the activity of the wild-type enzyme (Bjelke,
J. R., Christensen, J., Branner, S., Wagtmann, N., Olsen, C.
Kanstrup, A. B., and Rasmussen, H. B. (2004). Tyrosine 547 con-
stitutes an essential part of the catalytic mechanism of dipeptidyl
peptidase IV. /. Biol Chem. 279:34691-34697). Is this tyrosine
required for activity of DDP-IV? Why does the replacement of a
tyrosine with a phenylalanine abolish the enzyme activity?

10 . Acetylcholinesterase (AChE) catalyzes the breakdown of the neu-
rotransmitter acetylcholine to acetate and choline. This enzyme
contains a catalytic triad with the residues His, Glu, and Ser. The
catalytic triad enhances the nucleophilicity of the serine residue.
The nucleophilic oxygen of serine attacks the carbonyl carbon of
acetylcholine to form a tetrahedral intermediate.

.A,

H 3 CA ^Cr

Acetylcholine

(CH 2 ) 2 + h 2 o

N©(CH 3 ) 3

AChE ?

h 3 c^coo 0

^(CH 2 ) 2

HO— CH 2 ""N©

The nerve agent sarin is an extremely potent inactivator of AChE.
Sarin is an irreversible inhibitor that covalently modifies the ser-
ine residue in the active site of AChE.

/ \

O OCH

Sarin

(a) Diagram the expected arrangement of the amino acids in the
catalytic triad.

(b) Propose a mechanism for the covalent modification of AChE
by sarin.

194 CHAPTER 6 Mechanisms of Enzymes

11. Catalytic antibodies are potential therapeutic agents for drug
overdose and addiction. For example, a catalytic antibody that
catalyzes the breakdown of cocaine before it reached the brain
would be an effective detoxification treatment for drug abuse and
addiction. The phosphonate analog below was used to raise an
anticocaine antibody that catalyzes the rapid hydrolysis of co-
caine. Explain why this phosphonate ester was chosen to produce
a catalytic antibody.

Phosphonate analog

(-) - Cocaine

Ecgonine Benzoic acid

methyl ester

12. In the chronic lung disease emphysema, the lung s air sacs (alve-
oli), where oxygen from the air is exchanged for carbon dioxide in
the blood, degenerate. a \ -Proteinase inhibitor deficiency is a
genetic condition that runs in certain families and results from
mutations in critical amino acids in the sequence of a 1 -proteinase
inhibitor. The individuals with mutations are more likely to de-
velop emphysema, a 1 -Proteinase inhibitor is produced by the
liver and then circulates in the blood, al -Proteinase inhibitor is a
protein that serves as the major inhibitor of neutrophil elastase,
a serine protease present in the lung. Neutrophil elastase cleaves
the protein elastin, which is an important component for lung
function. The increased rate of elastin breakdown in lung tissue is
believed to cause emphysema. One treatment for a 1 -proteinase
inhibitor deficiency is to give the patient human wild-type
a 1 -proteinase inhibitor (derived from large pools of human
plasma) intravenously by injecting the protein directly into the
bloodstream.

(a) Explain the rational for the treatment with wild-type
a 1 -proteinase inhibitor.

(b) This treatment involves the intravenous administration
of the wild- type a 1 -proteinase inhibitor. Explain why
a 1 -proteinase inhibitor cannot be taken orally.

Selected Readings

General

Fersht, A. (1985). Enzyme Structure and Mechanism ,
2nd ed. (New York: W. H. Freeman).

Binding and Catalysis

Bartlett, G. J., Porter, C. T., Borkakoti, N. and
Thornton, J. M. (2002). Analysis of catalytic
residues in enzyme active sites. /. Mol. Biol.
324:105-121.

Bruice, T. C. and Pandrit, U. K. (1960). Intramole-
cular models depicting the kinetic importance of
“fit” in enzymatic catalysis. Proc. Natl. Acad. Sci.
USA. 46:402-404.

Hackney, D. D. (1990). Binding energy and catalysis.
In The Enzymes , Vol. 19, 3rd ed., D. S. Sigman and P.
D. Boyer, eds. (San Diego: Academic Press), pp. 1-36.

Jencks, W. P. (1987). Economics of enzyme catalysis.
Cold Spring Harbor Symp. Quant. Biol. 52:65-73.

Kraut, J. (1988). How do enzymes work? Science
242:533-540.

Neet, K. E. (1998). Enzyme catalytic power mini-
review series./. Biol. Chem. 273:25527-25528, and
related papers on pages 25529-25532, 26257-26260,
and 27035-27038.

Pauling, L. (1948) Nature of forces between large
molecules of biological interest. Nature
161:707-709.

Schiott, B., Iversen, B. B., Madsen, G. K. H., Larsen,
F. K., and Bruice, T. C. (1998). On the electronic
nature of low-barrier hydrogen bonds in
enzymatic reactions. Proc. Natl. Acad. Sci. USA
95:12799-12802.

Shan, S.-U., and Herschlag, D. (1996). The change
in hydrogen bond strength accompanying charge
rearrangement: implications for enzymatic cataly-
sis. Proc. Natl. Acad. Sci. USA 93:14474-14479.

Transition-State Analogs

Schramm, V. L. (1998). Enzymatic transition states
and transition state analog design. Annu. Rev.
Biochem. 67:693-720.

Wolfenden, R., and Radzicka, A. (1991). Transi-
tion-state analogues. Curr. Opin. Struct. Biol.
1:780-787.

Specific Enzymes

Cassidy, C. S., Lin, J., and Frey, P. A. (1997). A new
concept for the mechanism of action of chymo-
typsin: the role of the low-barrier hydrogen bond.
Biochem. 36:4576-4584.

Blacklow, S. C., Raines, R. T., Lim, W. A., Zamore,
P. D., and Lnowles, J. R. (1988). Triosephosphate
isomerase catalysis is diffusion controlled.
Biochem. 27:1158-1167.

Selected Readings 195

Davies, G. J., Mackenzie, L., Varrot, A., Dauter, M.,
Brzozowski, A. M., Schiilein, M., and Withers, S. G.
(1998). Snapshots along an enzymatic reaction
coordinate: analysis of a retaining (3 -glycoside
hydrolase. Biochem. 37:11707-11713.

Dodson, G., and Wlodawer, A. (1998). Catalytic
triads and their relatives. Trends Biochem. Sci.
23:347-352.

Frey, P. A., Whitt, S. A., and Tobin, J. B. (1994). A
low-barrier hydrogen bond in the catalytic triad of
serine proteases. Science. 264:1927-1930.

Getzoff, E. D., Cabelli, D. E., Fisher, C. L., Parge,

H. E., Viezzoli, M. S., Banci, L., and Hallewell, R. A.
(1992). Faster superoxide dismutase mutants de-
signed by enhancing electrostatic guidance. Nature.
358:347-351.

Harris, T. K., Abeygunawardana, C., and Mildvan,
A. S. (1997). NMR studies of the role of hydrogen
bonding in the mechanism of triosephosphate iso-
merase. Biochem. 36:14661-14675.

Huber, R., and Bode, W. (1978). Structural basis of
the activation and action of trypsin. Ace. Chem. Res.
11:114-122.

Kinoshita, T., Nishio, N., Nakanishi, I., Sato, A.,
and Fujii, T. (2003). Structure of bovine adeno-
sine deaminase complexed with 6-hydroxy- 1,6-
dihydropurine riboside. Acta Cryst. D59:299-303.

Kirby, A. J. (2001). The lysozyme mechanism sorted —
after 50 years. Nature Struct. Biol. 8:737-739.

Knolwes, J. R. (1991) Enzyme catalysis: not differ-
ent, just better. Nature. 350:121-124.

Knowles, J. R., and Albery, W. J. (1977). Perfection
in enzyme catalysis: the energetics of triosephos-
phate isomerase. Ace. Chem. Res. 10:105-111.

Kuser, P., Cupri, F., Bleicher, L., and Polikarpov, I.
(2008). Crystal structure of yeast hexokinase PI in
complex with glucose: a classical “induced fit” ex-
ample revisited. Proteins. 72:731-740.

Lin, J., Cassidy, C. S., and Frey, P. A. (1998). Corre-
lations of the basicity of His-57 with transition
state analogue binding, substrate reactivity, and
the strength of the low-barrier hydrogen bond in
chymotrypsin. Biochem. 37:11940-11948.

Lodi, P. J., and Knowles, J. R. (1991). Neutral
imidazole is the electrophile in the reaction cat-
alyzed by triosephosphate isomerase: structural
origins and catalytic implications. Biochem.
30:6948-6956.

Parthasarathy, S., Ravinda, G., Balaram, H.,
Balaram, P., and Murthy, M. R. N. (2002). Struc-
ture of the plasmodium falciparum triosephos-
phate isomerase — phosphoglycolate complex in
two crystal forms: characterization of catalytic
open and closed conformations in the ligand-
bound state. Biochem. 41:13178-13188.

Paetzel, M., and Dalbey, R. E. (1997). Catalytic
hydroxyl/amine dyads within serine proteases.
Trends Biochem. Sci. 22:28-31.

Perona, J. J., and Craik, C. S. (1997). Evolutionary
divergence of substrate specificity within the
chymotrypsin-like serine protease fold. /. Biol.
Chem. 272:29987-29990.

Schafer T., Borchert T. W., Nielsen V. S., Skager-
lind P., Gibson K., Wenger K., Hatzack F., Nilsson
L. D., Salmon S., Pedersen S., Heldt-Hansen H. P.,
Poulsen P. B., Lund H., Oxenboll K. M., Wu,

G. F., Pedersen H. H., Xu, H. (2007). Industrial
enzymes. Adv. Biochem. Eng. Biotechnol. 2007
105:59-131.

Steitz, T. A., and Shulman, R. G. (1982). Crystallo-
graphic and NMR studies of the serine proteases.
Annu. Rev. Biophys. Bioeng. 11:419-444.

Von Dreele, R. B. (2005). Binding of N-acetylglu-
cosamine oligosaccharides to hen egg-white
lysozyme: a powder diffraction study. Acta
Crystallographic. D6 1:22-32.

Zhou, G., Somasundaram, T., Blanc, E., Parthasarathy,
G., Ellington, W. R., and Chapman, M. S. (1998).
Transition state structure of arginine kinase: im-
plications for catalysis of bimolecular reactions.
Proc. Natl. Acad. Sci. USA 95:8449-8454.

o c

_ o

° o o o

° o

o o

° c

o o

Coenzymes and Vitamins

E volution has produced a spectacular array of protein catalysts but the catalytic
repertoire of an organism is not limited by the reactivity of amino acid side
chains. Other chemical species, called cofactors, often participate in catalysis.
Cofactors are required by inactive apoenzymes (proteins only) to convert them to active
holoenzymes. There are two types of cofactors: essential ions (mostly metal ions) and
organic compounds known as coenzymes (Figure 7.1). Both inorganic and organic co-
factors become essential portions of the active sites of certain enzymes.

Many of the minerals required by all organisms are essential because they are cofac-
tors. Some essential ions, called activator ions , are reversibly bound and often participate
in the binding of substrates. In contrast, some cations are tightly bound and frequently
participate directly in catalytic reactions.

Coenzymes act as group -transfer reagents. They accept and donate specific chemi-
cal groups. For some coenzymes, the group is simply hydrogen or an electron but other
coenzymes carry larger, covalently attached chemical groups. These mobile metabolic
groups are attached at the reactive center of the coenzyme. (Either the mobile metabolic
group or the reactive center is shown in red in the structures presented in this chapter.)
We can simplify our study of coenzymes by focusing on the chemical properties of their
reactive centers. The two classes of coenzymes are described in Section 7.2.

We begin this chapter with a discussion of essentialion cofactors. Much of the rest
of the chapter is devoted to the more complex organic cofactors. In mammals, many of
these coenzymes are derived from dietary precursors called vitamins. We therefore dis-
cuss vitamins in this chapter. We conclude with a look at a few proteins that are coen-
zymes. Most of the structures and reactions presented here will be encountered in later
chapters when we discuss particular metabolic pathways.

Cofactors

Essential ions Coenzymes

Activator ions Metal ions of Cosubstrates Prosthetic groups

(loosely bound) metalloenzymes (loosely bound) (tightly bound)
(tightly bound)

Finally, we come to a group of com-
pounds which have only been known
for a relatively short time ; but which
during this short time have attracted
very considerable attention ; both from
chemists and from the public at
large. Who today is unacquainted
with vitamins, these mysterious sub-
stances which are of such immense
significance for life, vita, itself and
which have thus justifiably taken
their name from it?

— H.G. Soderbaum Presentation
speech for the Nobel Prize in
chemistry to Adolf Windaus, 1 928

◄ Figure 7.1

Types of cofactors. Essential ions and coen-
zymes can be further distinguished by the
strength of interaction with their apoenzymes.

Top: Nicotinamide adenine dinucleotide (NAD®), a coenzyme derived from the vitamin nicotinic acid (niacin). NAD® is
an oxidizing agent.

196

7.2 Coenzyme Classification 197

7.1 Many Enzymes Require Inorganic Cations

Over a quarter of all known enzymes require metallic cations to achieve full catalytic ac-
tivity. These enzymes can be divided into two groups: metal-activated enzymes and
metalloenzymes. Metal-activated enzymes either have an absolute requirement for added
metal ions or are stimulated by the addition of metal ions. Some of these enzymes re-
quire monovalent cations such as K® and others require divalent cations such as Ca® or
Mg®. Kinases, for example, require magnesium ions for the magnesium- ATP complex
they use as a phosphoryl group donating substrate. Magnesium shields the negatively
charged phosphate groups of ATP making them more susceptible to nucleophilic attack
(Section 10.6).

Metalloenzymes contain firmly bound metal ions at their active sites. The ions
most commonly found in metalloenzymes are the transition metals, iron and zinc, and
less often, copper and cobalt. Metal ions that bind tightly to enzymes are usually re-
quired for catalysis. The cations of some metalloenzymes can act as electrophilic cata-
lysts by polarizing bonds. For example, the cofactor for the enzyme carbonic anhydrase
is an electrophilic zinc atom bound to the side chains of three histidine residues and to
a molecule of water. Binding to Zn® causes the water to ionize more readily. A basic
carboxylate group of the enzyme removes a proton from the bound water molecule,
producing a nucleophilic hydroxide ion that attacks the substrate (Figure 7.2). This en-
zyme has a very high catalytic rate partly because of the simplicity of its mechanism
(Section 6.4). Many other zinc metalloenzymes activate bound water molecules in this
fashion.

The ions of other metalloenzymes can undergo reversible oxidation and reduction
by transferring electrons from a reduced substrate to an oxidized substrate. For example,
iron is part of the heme group of catalase, an enzyme that catalyzes the degradation of
H 2 0 2 . Similar heme groups also occur in cytochromes, electron-transferring proteins
found associated with specific metalloenzymes in mitochondria and chloroplasts. Non-
heme iron is often found in metalloenzymes in the form of iron-sulfur clusters (Figure 7.3).
The most common iron-sulfur clusters are the [2 Fe-2 S] and [4 Fe-4 S] clusters in
which the iron atoms are complexed with an equal number of sulfide ions from H 2 S
and — S® groups from cysteine residues. Iron-sulfur clusters mediate some oxidation-
reduction reactions. Each cluster, whether it contains two or four iron atoms, can accept
only one electron in an oxidation reaction.

7.2 Coenzyme Classification

Coenzymes can be classified into two types based on how they interact with the apoen-
zyme (Figure 7.1). Coenzymes of one type — often called cosubstrates — are actually sub-
strates in enzyme-catalyzed reactions. A cosubstrate is altered in the course of the reac-
tion and dissociates from the active site. The original structure of the cosubstrate is
regenerated in a subsequent reaction catalyzed by another enzyme. The cosubstrate is
recycled repeatedly within the cell, unlike an ordinary substrate whose product typically
undergoes further transformation. Cosubstrates shuttle mobile metabolic groups
among different enzyme -catalyzed reactions.

The second type of coenzyme is called a prosthetic group. A prosthetic group re-
mains bound to the enzyme during the course of the reaction. In some cases the pros-
thetic group is covalently attached to its apoenzyme, while in other cases it is tightly
bound to the active site by many weak interactions. Like the ionic amino acid residues
of the active site, a prosthetic group must return to its original form during each full
catalytic event or the holoenzyme will not remain catalytically active. Cosubstrates and
prosthetic groups are part of the active site of enzymes. They supply reactive groups
that are not available on the side chains of amino acid residues.

Every living species uses coenzymes in a diverse number of important enzyme-
catalyzed reactions. Most of these species are capable of synthesizing their coenzymes
from simple precursors. This is especially true in four of the five kingdoms — prokary-
otes, protists, fungi, and plants — but animals have lost the ability to synthesize some

Refer to Figure 1.1 for a table of the
essential elements.

co 2 -^ ^co 2

h 2 o^ ^h 2 o

▲ Figure 7.2

Mechanism of carbonic anhydrase. The zinc
ion in the active site promotes the ionization
of a bound water molecule. The resulting
hydroxide ion attacks the carbon atom of
carbon dioxide, producing bicarbonate,
which is released from the enzyme.

Review Section 4.12 for the structure
of heme.

Cytochromes will be discussed in
Section 7.16.

198 CHAPTER 7 Coenzymes and Vitamins

▲ Figure 7.3

Iron-sulfur clusters. In each type of iron-
sulfur cluster, the iron atoms are complexed
with an equal number of sulfide ions (S 2- )
and with the thiolate groups of the side
chains of cysteine residues.

Table 7.1 Some vitamins and their

associated deficiency diseases

Vitamin

Disease

Ascorbate (C)

Scurvy

Thiamine (B-|)

Beriberi

Riboflavin (B 2 )

Growth retardation

Nicotinic acid (B 3 ) Pellagra

Pantothenate (B 5 )

Dermatitis in chickens

Pyridoxal (B 6 )

Dermatitis in rats

Biotin (B 7 )
Folate (B 9 )

Dermatitis in humans
Anemia

Cobalamin (B 12 )

Pernicious anemia

The structure and chemistry of
nucleotides is discussed in more detail
in Chapter 19.

coenzymes. Mammals (including humans) need a source of coenzymes in order to sur-
vive. The ones they cant synthesize are supplied by nutrients, usually in small amounts
(micrograms or milligrams per day). These essential compounds are called vitamins and
animals rely on other organisms to supply these micronutrients. The ultimate sources
of vitamins are usually plants and microorganisms. Most vitamins are coenzyme
precursors — they must be enzymatically transformed to their corresponding coenzymes.

A vitamin-deficiency disease can result when a vitamin is deficient or absent in the
diet of an animal. Such diseases can be overcome or prevented by consuming the appro-
priate vitamin. Table 7. 1 lists nine vitamins and the diseases associated with their defi-
ciencies. Each of these vitamins and their metabolic roles are discussed below. Most of
them are converted to coenzymes, sometimes after a reaction with ATP.

The word vitamin (originally spelled “vitamine”) was coined by Casimir Funk in
1912 to describe a “vital amine” from brown rice that cured beriberi, a nutritional-defi-
ciency disease that results in neural degeneration. The term vitamin has been retained
even though many vitamins proved not to be amines. Beriberi was first described in
birds and then in humans whose diets consisted largely of polished rice. Christiaan Eijk-
man, a Dutch physician working in what was then the Dutch East Indies (now Indone-
sia), was the first to notice that chickens fed polished rice leftover from the local hospital
developed beriberi but they recovered when they were fed brown rice. This discovery
led eventually to isolation of an antiberiberi substance from the skin that covers brown
rice. This substance became known as vitamin B x (thiamine).

Two broad classes of vitamins have since been identified: water-soluble (such as B
vitamins) and fat-soluble (also called lipid vitamins). Water-soluble vitamins are
required daily in small amounts because they are readily excreted in the urine and the
cellular stores of their coenzymes are not stable. Conversely, lipid vitamins such as vita-
mins A, D, E, and K, are stored by animals and excessive intakes can result in toxic con-
ditions known as hypervitaminoses. It’s important to note that not all vitamins are
coenzymes or their precursors (see Box 7.4 and Section 7.14).

The most common coenzymes are listed in Table 7.2 along with their metabolic
role and their vitamin source. The following sections describe the structures and func-
tions of these common coenzymes.

7.3 ATP and Other Nucleotide Cosubstrates

A number of nucleosides and nucleotides are coenzymes. Adenosine triphosphate (ATP) is
by far the most abundant. Other common examples are GTP, S-adenosylmethionine, and
nucleotide sugars such as uridine diphosphate glucose (UDP-glucose). ATP (Figure 7.4)
is a versatile reactant that can donate its phosphoryl, pyrophosphoryl, adenylyl (AMP),
or adenosyl groups in group -transfer reactions.

The most common reaction involving ATP is phosphoryl group transfer. In reac-
tions catalyzed by kinases, for example, the y-phosphoryl group of ATP is transferred to
a nucleophile leaving ADP. The second most common reaction is nucleotidyl group
transfer (transfer of the AMP moiety) leaving pyrophosphate (PPj). ATP plays a central
role in metabolism. Its role as a “high energy” cofactor is described in more detail in
Chapter 10, “Introduction to Metabolism.”

ATP is also the source of several other metabolite coenzymes. One, S-adenosylme-
thionine (Figure 7.5), is synthesized by the reaction of methionine with ATP.

Methionine + ATP > S-Adenosylmethionine + Pj + PPj (7.1)

The normal thiomethyl group of methionine ( — S — CH 3 ) is not very reactive but the posi-
tively charged sulfonium of 5 - adenosylmethionine is highly reactive. S-adenosylmethionine

◄ Brown rice and white rice. Brown rice (top left) has been processed to remove the outer husks but it
retains part of the outer skin or “bran.” This skin contains thiamine (vitamin B^. Further processing
of the grain yields white rice (middle left), which lacks thiamine.

7.3 ATP and Other Nucleotide Cosubstrates 199

Table 7.2 Major coenzymes

Coenzyme

Vitamin source

Major metabolic roles

Mechanistic role

Adenosine triphosphate (ATP)

—

Transfer of phosphoryl or
nucleotidyl groups

Cosubstrate

S-Adenosylmethionine

—

Transfer of methyl groups

Cosubstrate

Uridine diphosphate glucose

—

Transfer of glycosyl groups

Cosubstrate

Nicotinamide adenine dinucleotide (NAD®)
and nicotinamide adenine dinucleotide
phosphate (NADP®)

Niacin (B 3 )

Oxidation-reduction reactions
involving two-electron transfer

Cosubstrate

Flavin mononucleotide (FMN) and flavin
adenine dinucleotide (FAD)

Riboflavin (B 2 )

Oxidation-reduction reactions involving
one- and two-electron transfers

Prosthetic group

Coenzyme A (CoA)

Pantothenate (B 5 )

Transfer of acyl groups

Cosubstrate

Thiamine pyrophosphate (TPP)

Thiamine (B^

Transfer of multi-carbon fragments contain-
ing a carbonyl group

Prosthetic group

Pyridoxal phosphate (PLP)

Pyridoxine (B 6 )

Transfer of groups to and from amino acids

Prosthetic group

Biotin

Biotin (B 7 )

ATP-dependent carboxylation of substrates or
carboxyl-group transfer between substrates

Prosthetic group

Tetrahydrofolate

Folate

Transfer of one-carbon substituents, especially
formyl and hydroxymethyl groups; provides
the methyl group for thymine in DNA

Cosubstrate

Cobalamin

Cobalamin (B 12 )

Intramolecular rearrangements,
transfer of methyl groups.

Prosthetic group

Lipoamide

—

Oxidation of a hydroxyalkyl group from TPP
and subsequent transfer as an acyl group

Prosthetic group

Retinal

Vitamin A

Vision

Prosthetic group

Vitamin K

Carboxylation of some glutamate residues

Prosthetic group

Ubiquinone (Q)

—

Lipid-soluble electron carrier

Cosubstrate

Heme Group

—

Electron transfer

Prosthetic group

reacts readily with nucleophilic acceptors and is the donor of almost all the methyl y^g thermodynamics of reactions involv-

groups used in biosynthetic reactions. For example, it is required for conversion of the j n g ^yp j s explained in Section 10.6.

hormone norepinephrine to epinephrine.

Norepinephrine

Epinephrine (7.2)

O O

0 O— P— O— P —

.©

▲ Figure 7.4

ATP. The nitrogenous base adenine is linked to a ribose bearing three phosphoryl groups. Transfer of
a phosphoryl group (red) generates ADP, and transfer of a nucleotidyl group (AMP, blue) generates
pyrophosphate.

▲ Figure 7.5

S-Adenosylmethionine. The activated methyl
group of this coenzyme is shown in red.

200 CHAPTER 7 Coenzymes and Vitamins

BOX 7.1 MISSING VITAMINS

Whatever happened to vitamin B 4 and vitamin B 8 ? They are
never listed in the textbooks but you’ll often find them sold
in stores that cater to the demand for supplements that might
make you feel better and live longer.

Vitamin B 4 was adenine, the base found in DNA and
RNA. We now know that it’s not a vitamin. All species, in-
cluding humans, can make copious quantities of adenine
whenever it’s needed (Sections 18.1 and 18.2). Vitamin B 8
was inositol, a precursor of several important lipids
(Figure 8.16 and Section 9.12C). It’s no longer considered a
vitamin.

If you know anyone who is paying money for vitamin B 4
and B 8 supplements then here’s your chance to be helpful.
Tell them why they’re wasting their money.

▲ P.T. Barnum. P.T. Barnum was a famous American showman.
He’s credited with saying, “There’s a sucker born every minute.”
It’s likely that the memorable phrase was coined by one of his
rivals and later attributed to Barnum in order to discredit him.

Methylation reactions that require S-adenosylmethionine include methylation of phos-
pholipids, proteins, DNA, and RNA. In plants, S-adenosylmethionine — as a precursor
of the plant hormone ethylene — is involved in regulating the ripening of fruit.

Nucleotide-sugar coenzymes are involved in carbohydrate metabolism. The most
common nucleotide sugar, uridine diphosphate glucose (UDP-glucose), is formed by
the reaction of glucose 1 -phosphate with uridine triphosphate (UTP) (Figure 7.6 ).
UDP-glucose can donate its glycosyl group (shown in red) to a suitable acceptor, releas-
ing UDP. UDP-glucose is regenerated when UDP accepts a phosphoryl group from ATP
and the resulting UTP reacts with another molecule of glucose 1 -phosphate.

Both the sugar and the nucleoside of nucleotide-sugar coenzymes may vary. Later
on, we will encounter CDP, GDP, and ADP variants of this coenzyme.

7.4 NAD© and NADP©

The nicotinamide coenzymes are nicotinamide adenine dinucleotide (NAD®) and the
closely related nicotinamide adenine dinucleotide phosphate (NADP®). These were the
first coenzymes to be recognized. Both contain nicotinamide, the amide of nicotinic
acid (Figure 7.7 ). Nicotinic acid (also called niacin) is the factor missing in the disease
pellagra. Nicotinic acid or nicotinamide is essential as a precursor of NAD® and
NADP®. (In many species, tryptophan is degraded to nicotinic acid. Dietary trypto-
phan can therefore spare some of the requirement for niacin or nicotinamide.)

The nicotinamide coenzymes play a role in many oxidation-reduction reactions.
They assist in the transfer of electrons to and from metabolites (Section 10.9). The oxi-
dized forms, NAD® and NADP®, are electron deficient and the reduced forms, NADH
and NADPH, carry an extra pair of electrons in the form of a covalently bound hydride
ion. The structures of these coenzymes are shown in Figure 7.8 . Both coenzymes con-
tain a phosphoanhydride linkage that joins two 5' -nucleotides: AMP and the ribonu-
cleotide of nicotinamide, called nicotinamide mononucleotide (NMN) (formed from
nicotinic acid). In the case of NADP®, a phosphoryl group is present on the 2 '-oxygen
atom of the adenylate moiety.

Note that the ® sign in NAD® simply indicates that the nitrogen atom carries a
positive charge. This does not mean that the entire molecule is a positively charged ion;
in fact, it is negatively charged due to the phosphates. A nitrogen atom normally has

7.4 NAD© and NADP© 201

a-D-Glucose 1 -phosphate

◄ Figure 7.6

Formation of UDP-glucose catalyzed by UDP-
glucose pyrophosphorylase. An oxygen of the
phosphate group of a-D-glucose 1-phosphate
attacks the a-phosphorus of UTP. The PPj
released is rapidly hydrolyzed to 2Pj by the
action of pyrophosphatase. This hydrolysis
helps drive the pyrophosphorylase-catalyzed
reaction toward completion. The mobile gly-
cosyl group of UDP-glucose is shown in red.

seven protons and seven electrons. The outer shell has five electrons that can participate
in bond formation. In the oxidized form of the coenzyme (NAD® and NADP®)
the nicotinamide nitrogen is missing one of its electrons. It has only four electrons in
the outer shell and those are shared with adjacent carbon atoms to form a total of four
covalent bonds. (Each bond has a pair of electrons so the outer shell of the nitrogen
atom is filled with eight shared electrons.) This is why we normally associate the posi-
tive charge with the ring nitrogen atom as shown in Figure 7.8. In fact, the charge is
distributed over the entire aromatic ring.

The reduced form of the nitrogen atom has its normal, full complement of elec-
trons. In particular, the nitrogen atom has five electrons in its outer shell. Two of these
electrons (represented by dots in Figure 7.8) are a free pair of electrons. The other three
electrons participate in three covalent bonds.

NAD® and NADP® almost always act as cosubstrates for dehydrogenases. Pyri-
dine nucleotide-dependent dehydrogenases catalyze the oxidation of their substrates by
transferring two electrons and a proton in the form of a hydride ion (H®) to C-4 of the
nicotinamide group of NAD® or NADP®. This generates the reduced form, NADH or
NADPH, where a new C — H bond has formed at C-4 (one pair of electrons) and the
electron previously associated with the ring double bond has delocalized to the ring ni-
trogen atom. Thus, oxidation by pyridine nucleotides (or reduction, the reverse reac-
tion) always occurs two electrons at a time.

NADH and NADPH are said to possess reducing power (i.e., they are biological
reducing agents). The stability of reduced pyridine nucleotides allows them to carry
their reducing power from one enzyme to another, a property not shared by flavin

COOH

Nicotinic acid
(Niacin)

^nh 2

Nicotinamide

▲ Figure 7.7

Nicotinic acid (niacin) and nicotinamide.

NADH and NADPH exhibit a peak of
ultraviolet absorbance at 340 nm due
to the dihydropyridine ring, whereas
NAD® and NADP® do not absorb light
at this wavelength. The appearance
and disappearance of absorbance at
340 nm are useful for measuring the
rates of oxidation and reduction reac-
tions if they involve NAD® or NADP®.
(see Box 10.1).

202

CHAPTER 7 Coenzymes and Vitamins

Oxidized form

Reduced form

Nicotinamide

mononucleotide

(NMN)

Adenosine

monophosphate

(AMP)

H O

H H O

NAD® (NADP®)

NADH (NADPH)

▲ Figure 7.8

Oxidized and reduced forms of NAD (and
NADP). The pyridine ring of NAD© is re-
duced by the addition of a hydride ion to C-4
when NAD© is converted to NADH (and when
NADP© is converted to NADPH). In NADP©,
the 2'-hydroxyl group of the sugar ring of
adenosine is phosphorylated. The reactive
center of these coenzymes is shown in red.

coenzymes (Section 7.5). Most reactions forming NADH and NADPH are catabolic re-
actions and the subsequent oxidation of NADH by the membrane- associated electron
transport system is coupled to the synthesis of ATP. Most NADPH is used as a reducing
agent in biosynthetic reactions. The concentration of NADH is about ten times higher
than that of NADPH.

Lactate dehydrogenase is an oxidoreductase that catalyzes the reversible oxidation
of lactate. The enzyme is a typical NAD-dependent dehydrogenase. A proton is released
from lactate when NAD® is reduced.

OH O

H 3 c — CH— COO© + NAD® H 3 C— C —COO© + NADH + H®

Lactate Pyruvate (7.3)

NADH is a cosubtrate, like ATP. When the reaction is complete, the structure of the co-
substrate is altered and the original form must be regenerated in a separate reaction. In
this example, NAD® is reduced to NADH and the reaction will soon reach equilibrium
unless NADH is used up in a separate reaction where NAD® is regenerated. We de-
scribe one example of how this is accomplished in Section 11. 3B.

Figure 7.9 shows how both the enzyme and the coenzyme participate in the oxida-
tion of lactate to pyruvate catalyzed by lactate dehydrogenase. In this mechanism, the
coenzyme accepts a hydride ion at C-4 in the nicotinamide group. This leads to a re-
arrangement of bonds in the ring as electrons are shuffled to the positively charged
nitrogen atom. The enzyme provides an acid-base catalyst and suitable binding sites for
both the coenzyme and the substrate. Note that two hydrogens are removed from lac-
tate to produce pyruvate (Equation 7.3). One of these hydrogens is transferred to
NAD® as a hydride ion carrying two electrons and the other is transferred to His- 195
as a proton. The second hydrogen is subsequently released as H® in order to regenerate
the base catalyst (His- 195). There are many examples of NAD-dependent reactions
where the reduction of NAD® is accompanied by release of a proton so its quite common
to see NADH + H® on one side of the equation.

7.4 NAD© and NADP© 203

◄ Figure 7.9

Mechanism of lactate dehydrogenase. His-195,
a base catalyst in the active site, abstracts
a proton from the C-2 hydroxyl group of lac-
tate, facilitating transfer of the hydride ion
(H©) from C-2 of the substrate to C-4
of the bound NAD©. Arg-171 forms an ion
pair with the carboxylate group of the sub-
strate. In the reverse reaction, H© is trans-
ferred from the reduced coenzyme, NADH,
to C-2 of the oxidized substrate, pyruvate.

BOX 7.2 NAD BINDING TO DEHYDROGENASES

In the 1970s, structures were determined for four NAD-
dependent dehydrogenases: lactate dehydrogenase, malate
dehydrogenase, alcohol dehydrogenase, and glyceraldehyde
3 -phosphate dehydrogenase. Each of these enzymes is
oligomeric, with a chain length of about 350 amino acid
residues. These chains all fold into two distinct domains —
one to bind the coenzyme and one to bind the specific sub-
strate. For each enzyme, the active site is in the cleft between
the two domains.

As structures of more dehydrogenases were determined,
several conformations of the coenzyme-binding motif were ob-
served. Many of them possess one or more similar NAD- or
NADP-binding structures consisting of a pair of papaf} units

known as the Rossman fold after Michael Rossman, who first
observed them in nucleotide-binding proteins (see figure). Each
of the Rossman fold motifs binds to one half of the NAD® din-
ucleotide. All of these enzymes bind the coenzyme in the same
orientation and in a similar extended conformation.

Although many different dehydrogenases contain the
Rossman fold motif, the rest of the structures may be very
different and the dehydrogenases may not share significant
sequence similarity. It’s possible that all Rossman fold-
containing enzymes descend from a common ancestor, but
its also possible that the motifs evolved independently in dif-
ferent dehydrogenases. That would be another example of
convergent evolution.

◄ NAD-binding region of some dehydrogenases.

(a) The coenzyme is bound in an extended
conformation through interaction with two
side-by-side motifs known as Rossman folds.
The extended protein motifs form a p sheet of
six parallel p strands. The arrow indicates the
site where the hydride ion is added to C-4 of
the nicotinamide group, (b) NADH bound to a
Rossmann fold motif in rat lactate dehydroge-
nase [PDB 3H3F].

[Adapted from Rossman et al. (1975). The Enzymes,
Vol. 11, Part A, 3rd ed., P. D., Boyer, ed. (New York:
Academic Press), pp. 61-102.]

204 CHAPTER 7 Coenzymes and Vitamins

▲ These yellow FADs are not flavins but
Fish Aggregating Devices. They are buoys
tethered to the sea floor in order to attract
fish. This one has been deployed by the gov-
ernment of New South Wales off the east
coast of Australia. The strong ocean current
is threatening to carry it off.

7.5 FAD and FMN

The coenzymes flavin adenine dinucleotide (FAD) and flavin mononucleotide (FMN)
are derived from riboflavin, or vitamin B 2 . Riboflavin is synthesized by bacteria, pro-
tists, fungi, plants, and some animals. Mammals obtain riboflavin from food. Riboflavin
consists of the five-carbon alcohol ribitol linked to the N-10 atom of a heterocyclic ring
system called isoalloxazine (Figure 7.10a). The riboflavin-derived coenzymes are shown in
Figure 7.1 lb. Like NAD® and NADP® , FAD contains AMP and a diphosphate linkage.

Many oxidoreductases require FAD or FMN as a prosthetic group. Such enzymes
are called flavoenzymes or flavoproteins. The prosthetic group is very tightly bound,
usually noncovalently. By binding the prosthetic groups tightly, the apoenzymes protect
the reduced forms from wasteful reoxidation.

FAD and FMN are reduced to FADH 2 and FMNH 2 by taking up a proton and two
electrons in the form of a hydride ion (Figure 7.11). The oxidized enzymes are bright
yellow as a result of the conjugated double-bond system of the isoalloxazine ring sys-
tem. The color is lost when the coenzymes are reduced to FMNH 2 and FADH 2 .

FMNH 2 and FADH 2 donate electrons either one or two at a time, unlike NADH
and NADPH that participate exclusively in two-electron transfers. A partially oxidized
compound, FAD Ft* or FMNH-, is formed when one electron is donated. These interme-
diates are relatively stable free radicals called semiquinones. The oxidation ofFADH 2
and FMNH 2 is often coupled to reduction of a metalloprotein containing Fe^ (in an
[Fe-S] cluster). Because an iron-sulfur cluster can accept only one electron, the reduced
flavin must be oxidized in two one-electron steps via the semiquinone intermediate.
The ability of FMN to couple two-electron transfers with one-electron transfers is im-
portant in many electron transfer systems.

Crystals of Old Yellow Enzyme, a typi-

cal fiavoprotein, are shown in the 7.6 Coenzyme A and Acyl Carrier Protein

introduction to Chapter 5. Many metabolic processes depend on coenzyme A (CoA, or HS-CoA) including the

oxidation of fuel molecules and the biosynthesis of some carbohydrates and lipids.
This coenzyme is involved in acyl- group-transfer reactions in which simple carboxylic
acids and fatty acids are the mobile metabolic groups. Coenzyme A has three major
components: a 2-mercaptoethylamine unit that bears a free — SH group, the vitamin
pantothenate (vitamin B 5 , an amide of ( 3 - alanine and pantoate), and an ADP moiety

Figure 7.10 ►

Riboflavin and its coenzymes. (a) Riboflavin.
Ribitol is linked to the isoalloxazine ring sys-
tem. (b) Flavin mononucleotide (FMN, black)
and flavin adenine dinucleotide (FAD, black
and blue). The reactive center is shown in red.

(a)

h 3 c

H,C

Isoalloxazine

ch 2

CHOH

ch 2 oh

Ribitol

CHOH

,©

©r

7.6 Coenzyme A and Acyl Carrier Protein 205

FMNH* or FADH-
(semiquinone form)

◄ Figure 7.1 1

Reduction and reoxidation of FMN or FAD. The

conjugated double bonds between N-l and
N-5 are reduced by addition of a hydride ion
and a proton to form FMNH 2 or FADH 2 , re-
spectively, the hydroquinone form of each
coenzyme. Oxidation occurs in two steps.

A single electron is removed by a one-
electron oxidizing agent, with loss of a pro-
ton, to form a relatively stable free-radical
intermediate. This semiquinone is then oxi-
dized by removal of a proton and an electron
to form fully oxidized FMN or FAD. These
reactions are reversible.

FMNH 2 or FADH 2
(hydroquinone form)

whose 3' -hydroxyl group is esterified with a third phosphate group (Figure 7.12a).
The reactive center of CoA is the — SH group. Acyl groups covalently attach to the
— SH group to form thioesters. A common example is acetyl CoA (Figure 7.13), where
the acyl group is an acetyl moiety. Acetyl CoA is a “high energy” compound due to the
thioester linkage (Section 19.8). Coenzyme A was originally named for its role as the

v Figure 7.12

Coenzyme A and acyl carrier protein (ACP).

(a) In coenzyme A, 2-mercaptoethylamine
is bound to the vitamin pantothenate, which
in turn is bound via a phosphoester linkage
to an ADP group that has an additional
3'-phosphate group. The reactive center is
the thiol group (red), (b) In acyl carrier
protein, the phosphopantetheine prosthetic
group, which consists of the 2-mercap-
toethylamine and pantothenate moieties of
coenzyme A, is esterified to a serine residue
of the protein.

(b)

HS — CH 2 — CH 2 — N — C — CH 2 — CH 2 — N — C — CH
H H |

CH 3 O

I II

C — CH 2 — O — P — O — CH 2 — CH Serine

1 e 1

ch 3

Phosphopantetheine prosthetic group

Protein

206 CHAPTER 7 Coenzymes and Vitamins

H 3 c — c — S — CoA
Acetyl CoA

▲ Figure 7.13
Acetyl CoA

acetylation coenzyme. We will see acetyl CoA frequently when we discuss the metabo-
lism of carbohydrates, fatty acids, and amino acids.

Phosphopantetheine, a phosphate ester containing the 2-mercaptoethylamine and
pantothenate moieties of coenzyme A, is the prosthetic group of a small protein (77
amino acid residues) known as the acyl carrier protein (ACP). The prosthetic group is
esterified to ACP via the side-chain oxygen of a serine residue (Figure 7.12b). The — SH
of the prosthetic group of ACP is acylated by intermediates in the biosynthesis of fatty
acids (Chapter 16).

The metabolic role of pyruvate decar-
boxylase will be encountered in
Section 1 1.3. Transketolases are dis-
cussed in Section 12.9. The role of
TDP as a coenzyme in pyruvate dehy-
drogenase is described in Section 13.2.

7.7 Thiamine Diphosphate

Thiamine (or vitamin BJ contains a pyrimidine ring and a positively charged thia-
zolium ring (Figure 7.14a). The coenzyme is thiamine diphosphate (TDP), also called
thiamine pyrophosphate (TPP) in the older literature (Figure 7.14b). TDP is synthe-
sized from thiamine by enzymatic transfer of a pyrophosphoryl group from ATP.

About half a dozen decarboxylases (carboxylases) are known to require TDP as a
coenzyme. For example, TDP is the prosthetic group of yeast pyruvate decarboxylase
whose mechanism is shown in Figure 7.15. TDP is also a coenzyme involved in the
oxidative decarboxylation of a-keto acids other than pyruvate. The first steps in those
reactions proceed by the mechanism shown in Figure 7.15. In addition, TDP is a pros-
thetic group for enzymes known as transketolases that catalyze transfer between sugar
molecules of two -carbon groups that contain a keto group.

Figure 7.14 ►

Thiamine diphosphate (TDP). (a) Thiamine
(vitamin Bi). (b) Thiamine diphosphate
(TDP). The thiazolium ring of the coenzyme
contains the reactive center (red).

(a)

Pyrimidine

H 3 C ch 2 — ch 2 — OH

CH? — N

Thiazolium

ring

Thiamine (vitamin

(b) O O

Thiamine diphosphate
(TDP)

7.8 Pyridoxal Phosphate 207

TDP

Ylid

H,C

R - N ^/
i3

Enz — B:

/ e
h3C \ (

q — q Pyruvate
O \ 0°

Enz — B — H

Hydroxyethylthiamine

pyrophosphate

(HETDP)

H,C

R — N x S

I ^

h 3 c — ch t o v

Vl>H

Enz — B:

■»

Hz»C — C v

Acetaldehyde

H,C

R — N x /S

H 3 c — c — OH

©-

Enz— B — H '

Ylid

Enz— B -pH

H,C

R-N^S

n //

H 3 c-c-c v ^

Enz — B:

H,C

R — S
C

119

H 3 c — c — OH

TDP

Enz — B:

◄ Figure 7.15

Mechanism of yeast pyruvate decarboxylase.

The positive charge of the thiazolium ring of
TDP attracts electrons, weakening the bond
between C-2 and hydrogen. This proton is pre-
sumably removed by a basic residue of the
enzyme. Ionization generates a dipolar car-
banion known as an ylid (a molecule with
opposite charges on adjacent atoms). The
negatively charged C-2 attacks the electron-
deficient carbonyl carbon of the substrate
pyruvate and the first product (C0 2 ) is re-
leased. Two carbons of pyruvate are now at-
tached to the thiazole ring as part of a reso-
nance-stabilized carbanion. In the following
step, protonation of the carbanion produces
hydroxyethylthiamine diphosphate (HETDP).
HETDP is cleaved, releasing acetaldehyde
(the second product) and regenerating the
ylid form of the enzyme-TDP complex. TDP
re-forms when the ylid is protonated by the
enzyme.

The thiazolium ring of the coenzyme contains the reactive center. C-2 of TDP has
unusual reactivity; it is acidic despite its extremely high p K a in aqueous solution. Similarly,
recent experiments indicate that the p K a value for the ionization of hydroxyethylthiamine
diphosphate (HETDP) (i.e., formation of the dipolar carbanion) is changed from 15 in
water to 6 at the active site of pyruvate decarboxylase. This increased acidity is attributed
to low polarity of the active site, which also accounts for the reactivity of TDP.

7.8 Pyridoxal Phosphate

The B 6 family of water-soluble vitamins consists of three closely related molecules that
differ only in the state of oxidation or amination of the carbon bound to position 4 of
the pyridine ring (Figure 7.16a). Vitamin B 6 — most often pyridoxal or pyridoxamine —
is widely available from plant and animal sources. Induced B 6 deficiencies in rats result
in dermatitis and various disorders related to protein metabolism but actual vitamin

▲ Thiamine diphosphate bound to pyruvate
dehydrogenase. The coenzyme is bound in
an extended conformation and the diphos-
phate group is chelated to a magnesium
ion (green). [PDB 1PYD]

208 CHAPTER 7 Coenzymes and Vitamins

Figure 7.16 ►

Bg vitamins and pyridoxal phosphate, (a) Vita-
mins of the B 6 family: pyridoxine, pyridoxal,
and pyridoxamine. (b) Pyridoxal 5'-phosphate
(PLP). The reactive center of PLP is the
aldehyde group (red).

Pyridoxal

NHo

Pyridoxamine

Pyridoxal 5'-phosphate (PLP)

Figure 7.17 ►

Binding of substrate to a PLP-dependent
enzyme. The Schiff base linking PLP to a
lysine residue of the enzyme is replaced by
reaction of the substrate molecule with PLP.
The transimination reaction passes through
a geminal-diamine intermediate, resulting
in a Schiff base composed of PLP and the
substrate.

Internal aldimine
(PLP-enzyme)

B 6 deficiencies in humans are rare. Enzymatic transfer of the y-phosphoryl group from
ATP forms the coenzyme pyridoxal 5 '-phosphate (PLP) once vitamin B 6 enters a cell
(Figure 7.16b).

Pyridoxal phosphate is the prosthetic group for many enzymes that catalyze a vari-
ety of reactions involving amino acids such as isomerizations, decarboxylations, and
side-chain eliminations or replacements. In PLP-dependent enzymes, the carbonyl
group of the prosthetic group is bound as a Schiff base (imine) to the £- amino group of
a lysine residue at the active site. (A Schiff base results from condensation of a primary
amine with an aldehyde or ketone.) The enzyme-coenzyme Schiff base, shown on the
left in Figure 7.17, is sometimes referred to as an internal aldimine. PLP is tightly bound
to the enzyme by many weak noncovalent interactions; the additional covalent linkage
of the internal aldimine helps prevent loss of the scarce coenzyme when the enzyme is
not functioning.

Lys

7.9 Vitamin C 209

▲ Figure 7.18

Mechanism of transaminases. An amino acid displaces lysine from the internal aldimine that links PLP to the enzyme, generating an external
aldimine. Subsequent steps lead to the transfer of the amino group to PLP yielding an a-keto acid, which dissociates, and PMP, which remains
bound to the enzyme. If another a-keto acid enters, each step proceeds in reverse. The amino group is transferred to the a-keto acid producing a
new amino acid and regenerating the original PLP form of the enzyme.

The initial step in all PLP-dependent enzymatic reactions with amino acids is the
linkage of PLP to the a-amino group of the amino acid (formation of an external
aldimine). When an amino acid binds to a PLP-enzyme, a transimination reaction takes
place (Figure 7.17). This transfer reaction proceeds via a geminal- diamine intermediate
rather than via formation of the free-aldehyde form of PLP. Note that the Schiff bases
contain a system of conjugated double bonds in the pyridine ring ending with the posi-
tive charge on N-l. Similar ring structures with positively charged nitrogen atoms are
present in NAD®. The prosthetic group serves as an electron sink during subsequent
steps in the reactions catalyzed by PLP-enzymes. Once an a- amino acid forms a Schiff
base with PLP, electron withdrawal toward N-l weakens the three bonds to the a-carbon.
In other words, the Schiff base with PLP stabilizes a carbanion formed when one of the
three groups attached to the a- carbon of the amino acid is removed. Which group is
lost depends on the chemical environment of the enzyme active site.

Removal of the a- amino group from amino acids is catalyzed by transaminases
that participate in both the biosynthesis and degradation of amino acids (Chapter 17).
Transamination is the most frequently encountered PLP-dependent reaction. The
mechanism involves formation of an external aldimine (Figure 17.17) followed by re-
lease of the a-keto acid. The amino group remains bound to PLP forming pyridoxamine
phosphate (PMP) (Figure 7.18). The next step in transaminase reactions is the reverse
of the reaction shown in Figure 7.18 using a different a-keto acid as a substrate.

7.9 Vitamin C

The simplest vitamin is the antiscurvy agent ascorbic acid (vitamin C). Scurvy is a dis-
ease whose symptoms include skin lesions, fragile blood vessels, loose teeth, and bleed-
ing gums. The link between scurvy and nutrition was recognized four centuries ago
when British navy physicians discovered that citrus juice in limes and lemons were a
remedy for scurvy in sailors whose diet lacked fresh fruits and vegetables. It was not
until 1919, however, that ascorbic acid was isolated and shown to be the essential di-
etary component supplied by citrus juices.

► Limeys is the story of Dr. James Lind and his attempt to promote citrus fruit as a cure for scurvy
in the 1700s.

A specific transaminase is described
in Section 17.2B.

The Conquest of

sc u RVY

DAVID I. HARVIE

■ '

210 CHAPTER 7 Coenzymes and Vitamins

Chromosome 8

p23.2

p22.8

p22

p21 .3

021.2

Pl2
pi 1.21

—

ql 1 .21
ql 1 .20

q 1 2. 1

■

q12.3

q13.2
ql 5.0
q21 .1 1

q21 .80

q21 .9

q22.1

q22.2

q22.3

q23.1

q23.3

q24.12

q24.20

q24.21

q24.22

q24.28

v_v

q24.3

▲ The human GULO pseudogene is located
on the short arm of chromosome 8.

-2H @ , -2e°

▲ Figure 7.19

Ascorbic acid (vitamin C) and its dehydro, oxidized form.

Dehydroascorbic acid

Back in the 18th century it was not easy to convince authorities that a simple solu-
tion like citrus fruit would solve the problem of scurvy because there were many com-
peting theories. The story of Dr. James Lind and his efforts to convince the British navy
is just one of many stories associated with vitamin C. It shows us that scientific evidence
is not all that’s required in order to make changes in the way we do things. Eventually,
British sailors began to eat lemons and limes on a regular basis when they were at sea.
Not only did this reduce the incidences of scurvy but it also gave rise to a famous nick-
name for British sailors. They were called “limeys” although lemons were much more
effective than limes.

Ascorbic acid is a lactone, an internal ester in which the C-l carboxylate group is
condensed with the C-4 hydroxyl group, forming a ring structure. We now know that
ascorbic acid is not a coenzyme but acts as a reducing agent in several different enzy-
matic reactions (Figure 7.19). The most important of these reactions is the hydroxyla-
tion of collagen (Section 4.12). Most mammals can synthesize ascorbic acid but guinea
pigs, bats, and some primates (including humans) lack this ability and must therefore
rely on dietary sources.

In most cases, we don’t know very much about how certain enzymes disappeared
from some species leading to a reliance on external sources for some essential metabo-
lites. Most of the presumed gene disruption events happened so far in the distant past
that few traces remain in modern genomes. The loss of ability to make vitamin C is an
exception to that rule and serves as an instructive example of evolution.

Ascorbic acid is synthesized from D-glucose in a five-step pathway involving four
enzymes (the last step is spontaneous). The last enzyme in the pathway is L-glucono-

CHO

H — C — OH

HO — C — H

Enz r HO-C-H

H — C — OH

CH 2 OH

COO

D-Glucose

D-Glucuronic

acid

CHO

D-Glucuronic
acid lactone

L-Ascorbic

Acid

CH 2 OH

ch 2 oh

L-Gulono-

lactone

2-Keto

L-Gulono-

lactone

ChLOH

Enzyme
4

L-Gulono-
gamma-lactone
oxidase (GULO)

▲ Figure 7.20

Biosynthesis of ascorbic acid (vitamin C).

L-ascorbic acid is synthesized from D-glu-
cose. The last enzymatic step is catalyzed by
L-glucono-gamma-lactone oxidase (GULO),
an enzyme that is missing in most primates.

7.10 Biotin 211

Rat GULO gene

I II

-I I

IV V

VII VIII

-ii

IX X

XI XII

Human GULO pseudogene

◄ Figure 7.21

Comparison of the intact rat GULO gene and the
human pseudogene. The human pseudogene
is missing the first six exons and exon 11.

In addition, there are many mutations in the
remaining exons that prevent them from pro-
ducing protein product.

gamma-lactone oxidase (GULO) (Figure 7.20). GULO (the enzyme) is not present in
primates of the haplorrhini family (monkeys and apes), but it is present in the strepsir-
rhini (lemurs, lorises etc.). These groups diverged about 80 million years ago. This led to
the prediction that the GULO gene would be absent or defective in the monkeys and
apes but intact in the other primates.

The prediction was confirmed with the discovery of a human GULO pseudogene
on chromosome 8 in a block of genes that contains an active GULO gene in other ani-
mals. A comparison of the human pseudogene and a functional rat gene reveals many
differences (Figure 7.21). The human pseudogene is missing the first six exons of the
normal gene plus exon 11. The pseduogene in other apes is also missing these exons in-
dicating that the ancestor of all apes had a similar defective GULO gene.

The original mutation that made the GULO gene inactive isn’t known. Once inac-
tivated, the pseudogene accumulated additional mutations that became fixed by
random genetic drift. We can assume that lack of ability to synthesize vitamin C was
not detrimental in these species because they obtained sufficient quantities in their
normal diet.

7.10 Biotin

Biotin is a prosthetic group for enzymes that catalyze carboxyl group transfer reactions
and ATP-dependent carboxylation reactions. Biotin is covalently linked to the active
site of its host enzyme by an amide bond to the £-amino group of a lysine residue
(Figure 7.22).

Biotin

“i r

Lysine

HNi bNH *

\ /

HC — CH O NH

/ \ II I

H 2 C x ^ch — ch 2 — ch 2 — ch 2 — ch 2 — c — N — ch 2 — ch 2 — ch 2 — ch 2 — CH

Enzyme-bound biotin

C=0

◄ Figure 7.22

Enzyme-bound biotin. The carboxylate group
of biotin is covalently bound via amide link-
age to the £-amino group of a lysine residue
(blue). The reactive center of the biotin
moiety is N-l (red).

The pyruvate carboxylase reaction demonstrates the role of biotin as a carrier of
carbon dioxide (Figure 7.23). In this ATP-dependent reaction, pyruvate, a three-carbon
acid, reacts with bicarbonate forming the four-carbon acid oxaloacetate. Enzyme-
bound biotin is the intermediate carrier of the mobile carboxyl metabolic group. The
pyruvate carboxylase reaction is an important C0 2 fixation reaction. It is required in
the gluconeogenesis pathway (Chapter 11).

Biotin was first identified as an essential factor for the growth of yeast. Biotin defi-
ciency is rare in humans or animals on normal diets because biotin is synthesized by
intestinal bacteria and is required only in very small amounts (micrograms per day). A
biotin deficiency can be induced, however, by ingesting raw egg whites that contain a
protein called avidin. Avidin binds tightly to biotin making it unavailable for absorption

212 CHAPTER 7 Coenzymes and Vitamins

Voe

Bicarbonate

coo°

Enol pyruvate |

C — 0°

Biotin Carboxybiotin

coo°

Oxaloacetate

C = 0

Biotin

▲ Figure 7.23

Reaction catalyzed by pyruvate carboxylase. First, biotin, bicarbonate, and ATP react to form carboxybiotin. The carboxybiotinyl-enzyme complex provides
a stable, activated form of CO 2 that can be transferred to pyruvate. Next, the enolate form of pyruvate attacks the carboxyl group of carboxybiotin,
forming oxaloacetate and regenerating biotin.

from the intestinal tract. Avidin is denatured when eggs are cooked and it loses its affin-
ity for biotin.

A variety of laboratory techniques take advantage of the high affinity of avidin for
biotin. For example, a substance to which biotin is covalently attached can be extracted
from a complex mixture by affinity chromatography (Section 3.6) on a column of im-
mobilized avidin. The association constant for biotin and avidin is about 10 15 M -1 —
one of the tightest binding interactions known in biochemistry (see Section 4.9).

BOX 7.3 ONE GENE: ONE ENZYME

George Beadle and Edward Tatum wanted to test the idea
that each gene encoded a single enzyme in a metabolic path-
way. It was back in the late 1930s and this correspondence,
which we now take for granted, was still a hypothesis. Re-
member, this was a time when it wasn’t even clear whether
genes were proteins or some other kind of chemical.

Beadle and Tatum chose the fungus Neurospora crassa
for their experiments. Neurospora grows on a well-defined
medium needing only sugar and biotin (vitamin B 7 ) as sup-
plements. They reasoned that by irradiating Neurospora
spores with X rays they could find mutants that would grow
on rich supplemented medium but not on the simple defined
medium. All they had to do next was identify the one supple-
ment that needed to be added to the minimal medium to
correct the defect. This would identify a gene for an enzyme
that synthesized the now- essential supplement.

The 299th mutant required vitamin B 6 and the 1085th
mutant required vitamin B x . The B 6 and B x biosynthesis
pathways were the first two pathways to be identified in this
set of experiments. Later on, they worked out the genes/en-
zymes used in the tryptophan pathway. The results were pub-
lished in 1941 and Beadle and Tatum received the Nobel
Prize in Physiology or Medicine in 1958.

▲ Neurospora crassa growing on defined medium in a test tube. The

strains on the right are producing orange carotenoid and the ones on
the left are nonproducing strains.

(Source: Courtesy of Manchester University, United Kingdom).

7.11 Tetrahydrofolate

213

7.11 Tetrahydrofolate

The vitamin folate was first isolated in the early 1940s from green leaves, liver, and yeast.
Folate has three main components: pterin (2-amino-4-oxopteridine), ap-aminobenzoic
acid moiety, and a glutamate residue. The structures of pterin and folate are shown in
Figures 7.24a and 7.24b. Humans require folate in their diets because we cannot synthe-
size the pterin-p-aminobenzoic acid intermediate (PABA) and we cannot add glutamate
to exogenous PABA.

The coenzyme forms of folate, known collectively as tetrahydrofolate, differ from
the vitamin in two respects: they are reduced compounds (5,6,7,8-tetrahydropterins),
and they are modified by the addition of glutamate residues bound to one another
through y- glutamyl amide linkages (Figure 7.24c). The anionic polyglutamyl moiety,
usually five to six residues long, participates in the binding of the coenzymes to en-
zymes. When using the term tetrahydrofolate , keep in mind that it refers to compounds
that have polyglutamate tails of varying lengths.

Tetrahydrofolate is formed from folate by adding hydrogen to positions 5, 6, 7, and
8 of the pterin ring system. Folate is reduced in two NADPH-dependent steps in a reac-
tion catalyzed by dihydrofolate reductase (DHFR).

NADPH + H @ NADPH + H @

Folate 7,8-Dihydrofolate 5,6,7,8-Tetrahydrofolate

(7.4)

The primary metabolic function of dihydrofolate reductase is the reduction of di-
hydrofolate produced during the formation of the methyl group of thymidylate
(dTMP) (Chapter 18). This reaction, which uses a derivative of tetrahydrofolate, is an
essential step in the biosynthesis of DNA. Because cell division cannot occur when DNA
synthesis is interrupted, dihydrofolate reductase has been extensively studied as a target
for chemotherapy in the treatment of cancer (Box 18.4). In most species, dihydrofolate
reductase is a relatively small monomeric enzyme that has evolved efficient binding sites
for the two large substrates (folate and NADPH) (Figure 6.12).

▼ Figure 7.24

Pterin, folate, and tetrahydrofolate. Pterin
(a) is part of folate (b), a molecule contain-
ing p-ami nobenzoate (red) and glutamate
(blue), (c) The polyglutamate forms of
tetrahydrofolate usually contain five or six
glutamate residues. The reactive centers
of the coenzyme, N-5 and N-10, are shown
in red.

Tetrahydrofolate (Tetrahydrofolyl polyglutamate)

214 CHAPTER 7 Coenzymes and Vitamins

Figure 7.25 ►

One-carbon derivatives of tetra hydrofolate.

The derivatives can be interconverted enzy-
matically by the routes shown. (R represents
the benzoyl polyglutamate portion of
tetrahydrofolate.)

H 9 N

C — N — R

5-Methyltetrahydrofolate 5 # 10-Methylenetetrahydrofolate

▲ Many fruits and vegetables contain adequate
supplies of folate. Yeast and liver products
are also excellent sources of folate.

H 7 N

CH — CH — CH q

OH OH

▲ Figure 7.26

5,6,7,8-Tetrahydrobiopterin. The hydrogen
atoms lost on oxidation are shown in red.

H,N

'f"CH 2

O /

u HC — N — R

5 # 10-Methenyltetrahydrofolate

1 0-Formyltetrahydrofolate

5,6,7,8-Tetrahydrofolate is required by enzymes that catalyze biochemical transfers
of several one-carbon units. The groups bound to tetrahydrofolate are methyl, methylene,
or formyl groups. Figure 7.25 shows the structures of several one-carbon derivatives of
tetrahydrofolate and the enzymatic interconversions that occur among them. The one-
carbon metabolic groups are covalently bound to the secondary amine N-5orN-10of
tetrahydrofolate, or to both in a ring form. 10-Formyltetrahydro folate is the donor of
formyl groups and 5, 10-methylenetetrahydro folate is the donor of hydroxymethyl
groups.

Another pterin coenzyme, 5,6,7,8-tetrahydrobiopterin, has a three-carbon side
chain at C-6 of the pterin moiety in place of the large side chain found in tetrahydrofo-
late (Figure 7.26). This coenzyme is not derived from a vitamin but is synthesized by
animals and other organisms. Tetrahydrobiopterin is the cofactor for several hydroxy-
lases and will be encountered as a reducing agent in the conversion of phenylalanine to
tyrosine (Chapter 17). It also is required by the enzyme that catalyzes the synthesis of
nitric oxide from arginine (Section 17.12).

The sale of vitamins and supplements is big business in developed nations. It’s
often difficult to decide whether an extra supply of vitamins is necessary for good health
because the scientific evidence is often missing or contradictory. Folate (vitamin B 9 )
deficiency is uncommon in normal, healthy adults and children in developed nations
but there are documented cases of folate deficiency in pregnant women. A lack of
tetrahydrofolate can lead to anemia and to severe defects in the developing fetus.
While there are many fruits and vegetables that contain folate, it’s a good idea for preg-
nant women to supplement their diet with folate in order to ensure their own health
and that of the baby.

7.12 Cobalamin 215

7.12 Cobalamin

Cobalamin (vitamin B 12 ) is the largest B vitamin and was the last to be isolated. The
structure of cobalamin (Figure 7.27a) includes a corrin ring system that resembles the
porphyrin ring system of heme (Figure 4.37). Note that cobalamin contains cobalt
rather than the iron found in heme. The abbreviated structure shown in Figure 7.27b
emphasizes the positions of two axial ligands bound to the cobalt, a benzimida-
zole ribonucleotide below the corrin ring and an R group above it. In the coenzyme
forms of cobalamin, the R group is either a methyl group (in methylcobalamin) or a
5'-deoxyadenosyl group (in adenosylcobalamin).

Cobalamin is synthesized by only a few microorganisms. It is required as a mi-
cronutrient by all animals and by some bacteria and algae. Humans obtain cobalamin
from foods of animal origin. A deficiency of cobalamin can lead to pernicious anemia, a
potentially fatal disease in which there is a decrease in the production of blood cells by
bone marrow. Pernicious anemia can also cause neurological disorders. Most victims of
pernicious anemia do not secrete a necessary glycoprotein (called intrinsic factor) from
the stomach mucosa. This protein specifically binds cobalamin and the complex is ab-
sorbed by cells of the small intestine. Impaired absorption of cobalamin is now treated
by regular injections of the vitamin.

The role of adenosylcobalamin reflects the reactivity of its C — Co bond. The coen-
zyme participates in several enzyme-catalyzed intramolecular rearrangements in which a
hydrogen atom and a second group, bound to adjacent carbon atoms within a substrate,
exchange places (Figure 7.28a). An example is the methylmalonyl-CoA mutase reaction
(Figure 7.28b) that is important in the metabolism of odd-chain fatty acids (Chapter 16)
and leads to the formation of succinyl CoA, an intermediate of the citric acid cycle.

Methylcobalamin participates in the transfer of methyl groups, as in the regenera-
tion of methionine from homocysteine in mammals.

▲ Dorothy Crowfoot Hodgkin (1910-1994).

Hodgkin received the Nobel Prize in 1964
for determining the structure of vitamin B 12
(cobalamin). The structure of insulin, shown
in the photograph, was published in 1969.

▲ Figure 7.27

Cobalamin (vitamin B 12 ) and its coenzymes, (a) Detailed structure of cobalamin showing the corrin ring system (black) and 5,6-dimethylbenzimidazole
ribonucleotide (blue). The metal coordinated by corrin is cobalt (red). The benzimidazole ribonucleotide is coordinated with the cobalt of the corrin ring
and is also bound via a phosphoester linkage to a side chain of the corrin ring system, (b) Abbreviated structure of cobalamin coenzymes. A benzimida-
zole ribonucleotide lies below the corrin ring, and an R group lies above the ring.

216 CHAPTER 7 Coenzymes and Vitamins

Figure 7.28 ►

Intramolecular rearrangements catalyzed
by adenosylcobalamin-dependent enzymes.

(a) Rearrangement in which a hydrogen
atom and a substituent on an adjacent carbon
atom exchange places, (b) Rearrangement
of methylmalonyl CoA to succinyl CoA,
catalyzed by methylmalonyl-CoA mutase.

▲ Intestinal bacteria. Normal, healthy hu-
mans harbor billions of bacteria in their in-
testines. There are at least several dozen
different species. The one shown here is
Helicobacter pylori, which causes stomach
ulcers when it invades the stomach. The
bacteria are sitting on the surface of the
intestine that has many projections for ab-
sorbing nutrients. Other common species
are Escherichia coli and various species of
Actinomyces and Streptococcus. These bac-
teria help break down ingested food and
they supply many of the essential vitamins
and amino acids that humans need, espe-
cially cobalamin.

Figure 7.29 ►

Lipoamide. Lipoic acid is bound in amide
linkage to the e-amino group of a lysine
residue (blue) of dihydrolipoamide acyltrans-
ferases. The dithiolane ring of the lipoyllysyl
groups is extended 1.5 nm from the
polypeptide backbone. The reactive center
of the coenzyme is shown in red.

(a)

b — c— ;

e — C — H

b— C — H

e — c— :

(b)

H 0

o 1 11

°ooc— c — c-

S-CoA

Methylmalonyl-CoA

OOC— c — H

mutase

H — C — H

Adenosylcobalamin

h — c— : — s-

1 II

Methylmalonyl CoA

H 0

Succinyl CoA

coo°

5-Methyltetrahydrofolate

coo 0

h 3 n — ch

Tetrahydrofolate

H 3 N — CH

cn 2

w >

ch 2

oh 2

Homocysteine

methyltransferase

ch 2

Flomocysteine

Methylcobalamin

s — ch 3

Methionine

(7.5)

In this reaction, the methyl group of 5-methyltetrahydrofolate is passed to a reactive,
reduced form of cobalamin to form methylcobalamin that can transfer the methyl
group to the thiol side chain of homocysteine.

7.13 Lipoamide

The lipoamide coenzyme is the protein-bound form of lipoic acid. Lipoic acid is some-
times described as a vitamin but animals appear to be able to synthesize it. It is required
by certain bacteria and protozoa for growth. Lipoic acid is an eight- carbon carboxylic
acid (octanoic acid) in which two hydrogen atoms, on C-6 and C-8, have been replaced
by sulfhydryl groups in disulfide linkage. Lipoic acid does not occur free — it is cova-
lently attached via an amide linkage through its carboxyl group to the e- amino group of
a lysine residue of a protein (Figure 7.29). This structure is found in dihydrolipoamide
acyltransferases that are components of the pyruvate dehydrogenase complex and
related enzymes.

Lipoamide carries acyl groups between active sites in multienzyme complexes. For
example, in the pyruvate dehydrogenase complex (Section 12.2), the disulfide ring of

Lipoyllysyl group

1.5 nm

O C =0

8/ C H2 6 ||

h 2 c ch — ch 2 — ch 2 — ch 2 — ch 2 — c— n— ch 2 — ch 2 — ch 2 — ch 2 — ch

\ /

s — S NH

Lipoamide

Lysine side chain

7.14 Lipid Vitamins

217

the lipoamide prosthetic group reacts with HETDP (Figure 7.15) binding its acetyl
group to the sulfur atom attached to C-8 of lipoamide and forming a thioester. The acyl
group is then transferred to the sulfur atom of a coenzyme A molecule generating the
reduced (dihydrolipoamide) form of the prosthetic group.

CH 2

X X

h 2 c ch— r

h 3 c — c — s

ch 2

X X

h 2 c ch

(7.6)

The final step catalyzed by the pyruvate dehydrogenase complex is the oxidation of
dihydrolipoamide. In this reaction, NADH is formed by the action of a flavoprotein
component of the complex. The actions of the multiple coenzymes of the pyruvate de-
hydrogenase complex show how coenzymes, by supplying reactive groups that augment
the catalytic versatility of proteins, are used to conserve both energy and carbon building
blocks.

7.14 Lipid Vitamins

The structures of the four lipid vitamins (A, D, E, and K) contain rings and long
aliphatic side chains. The lipid vitamins are highly hydrophobic although each possesses
at least one polar group. In humans and other mammals, ingested lipid vitamins are ab-
sorbed in the intestine by a process similar to the absorption of other lipid nutrients
(Section 16.1a). After digestion of any proteins that may bind them, they are carried to
the cellular interface of the intestine as micelles formed with bile salts. The study of
these hydrophobic molecules has presented several technical difficulties so research on
their mechanisms has progressed more slowly than that on their water-soluble counter-
parts. Lipid vitamins differ widely in their functions, as we will see below.

A. Vitamin A

Vitamin A, or retinol, is a 20-carbon lipid molecule obtained in the diet either directly or
indirectly from /?- carotene. Carrots and other yellow vegetables are rich in /3- carotene, a
40-carbon plant lipid whose enzymatic oxidative cleavage yields vitamin A (Figure 7.30).
Vitamin A exists in three forms that differ in the oxidation state of the terminal func-
tional group: the stable alcohol retinol, the aldehyde retinal, and retinoic acid. Their hy-
drophobic side chain is formed from repeated isoprene units (Section 9.6).

All three vitamin A derivatives have important biological functions. Retinoic acid is
a signal compound that binds to receptor proteins inside cells; the ligand-receptor

◄ Figure 7.30

Formation of vitamin A from /2-carotene.

Vitamin A
(retinol form)

CH 2 OH

218 CHAPTER 7 Coenzymes and Vitamins

Vitamin D 3
(Cholecalciferol)

1,25-Dihydroxycholecalciferol
▲ Figure 7.31

Vitamin D 3 (cholecalciferol) and 1,25-
dihydroxycholecalciferol. (Vitamin D 2 has an
additional methyl group at C-24 and a trans
double bond between C-22 and C-23.) 1,25-
Dihydroxycholecalciferol is produced from
vitamin D 3 by two separate hydroxylations.

complexes then bind to chromosomes and can regulate gene expression during cell
differentiation. The aldehyde retinal is a light-sensitive compound with an important
role in vision. Retinal is the prosthetic group of the protein rhodopsin; absorption of a
photon of light by retinal triggers a neural impulse.

B. Vitamin D

Vitamin D is the collective name for a group of related lipids. Vitamin D 3 (cholecalcif-
erol) is formed nonenzymatically in the skin from the steroid 7-dehydrocholesterol
when humans are exposed to sufficient sunlight. Vitamin D 2 , a compound related to
vitamin D 3 (D 2 has an additional methyl group), is the additive in fortified milk. The
active form of vitamin D 3 , 1,25-dihydroxycholecalciferol, is formed from vitamin D 3 by
two hydroxylation reactions (Figure 7.31 ); vitamin D 2 is similarly activated. The active
compounds are hormones that help control Ca® utilization in humans — vitamin D
regulates both intestinal absorption of calcium and its deposition in bones. In vitamin D-
deficiency diseases, such as rickets in children and osteomalacia in adults, bones are
weak because calcium phosphate does not properly crystallize on the collagen matrix of
the bones.

C. Vitamin E

Vitamin E, or a- tocopherol (Figure 7.32), is one of several closely related tocopherols,
compounds having a bicyclic oxygen-containing ring system with a hydrophobic side
chain. The phenol group of vitamin E can undergo oxidation to a stable free radical.
Vitamin E is believed to function as a reducing agent that scavenges oxygen and free
radicals. This antioxidant action may prevent damage to fatty acids in biological
membranes. A deficiency of vitamin E is rare but may lead to fragile red blood cells
and neurological damage. The deficiency is almost always caused by genetic defects in
absorption of fat molecules. There is currently no scientific evidence to support claims
that vitamin E supplements in the diet of normal, healthy individuals will improve
health.

Phylloquinone (vitamin K) are impor-
tant components of photosynthesis
reaction centers in bacteria, algae,
and plants.

D. Vitamin K

Vitamin K (phylloquinone) (Figure 7.32) is a lipid vitamin from plants that is required
for the synthesis of some of the proteins involved in blood coagulation. It is a coenzyme
for a mammalian carboxylase that catalyzes the conversion of specific glutamate
residues to y-carboxyglutamate residues (Equation 7.7). The reduced (hydroquinone)
form of vitamin K participates in the carboxylation as a reducing agent. Oxidized
vitamin K has to be regenerated in order to support further modifications of clotting
factors. This is accomplished by vitamin K reductase.

Vitamin E
(u-tocopherol)

Figure 7.32 ►

Structures of vitamin E and vitamin K.

7.15 Ubiquinone 219

▲ Vitamin D and the evolution of skin color. Black skin protects cells from damage by sunlight but it may inhibit formation of vitamin D. This isn’t a
problem in Nairobi, Kenya (left) but it might be in Stockholm, Sweden (right). One hypothesis for the evolution of skin color suggests that light-
colored skin evolved in northern climates in order to increase vitamin D production.

Glutamate residue

y-Carboxyglutamate residue

'WV |\|

Vitamin K reductase

(7.7)

When calcium binds to the y-carboxyglutamate residues of the coagulation pro-
teins, the proteins adhere to platelet surfaces where many steps of the coagulation
process take place.

7.15 Ubiquinone

Ubiquinone — also called coenzyme Q and therefore abbreviated a Q” — is a lipid-soluble
coenzyme synthesized by almost all species. Ubiquinone is a benzoquinone with four sub-
stituents, one of which is a long hydrophobic chain. This chain of 6 to 10 isoprenoid units
allows ubiquinone to dissolve in lipid membranes. In the membrane, ubiquinone trans-
ports electrons between enzyme complexes. Some bacteria use menaquinone instead of
ubiquinone (Figure 7.33 a). An analog of ubiquinone, plastoquinone (Figure 7.33b), serves
a similar function in photosynthetic electron transport in chloroplasts (Chapter 15).

Ubiquinone is a stronger oxidizing agent than either NAD® or the flavin coen-
zymes. Consequently, it can be reduced by NADH or FADH 2 . Like FMN and FAD,
ubiquinone can accept or donate two electrons one at a time because it has three oxidation
states: oxidized Q, a partially reduced semiquinone free radical, and fully reduced QH 2 ,
called ubiquinol (Figure 7.34 ). Coenzyme Q plays a major role in membrane-associated
electron transport. It is responsible for moving protons from one side of the membrane
to the other by a process known as the Q cycle. (Chapter 14). The resulting proton
gradient contributes to ATP synthesis.

220 CHAPTER 7 Coenzymes and Vitamins

BOX 7.4 RAT POISON

Warfarin is an effective rat poison that has been used for
many decades. It’s a competitive inhibitor of vitamin K reduc-
tase, the enzyme that regenerates the reduced form of vitamin
K (Equation 7.7). Blocking the formation of blood clotting
factors leads to death in the rodents by internal bleeding. Ro-
dents are very sensitive to inhibition of vitamin K reductase.

Later on it was discovered that low concentrations of
warfarin were effective in individuals who suffer from excessive
blood clotting. The drug was renamed (e.g., Coumadin®) for
use in humans since its association with rat poison had a
somewhat negative connotation.

Vitamin K analogs are widely used as anticoagulants in
patients who are prone to thrombosis where they can prevent
strokes and other embolisms. Like all medications, the dosage
must be carefully regulated and controlled in order to prevent
adverse effects, but in this case the dosage is even more critical.

Since the drugs only affect the synthesis of new clotting fac-
tors, they often take several days to have an effect.This is why
patients will often be started at low dosages of these analogs
and the amount of drug will be increased slowly over the
course of many months.

▲ Warfarin. a A rat [Rattus norvegicus).

Figure 7.33 ►

Structures of (a)
menaquinone and (b) plasto-
quinone. The hydrophobic
tail of each molecule is
composed of 6 to 10 five-
carbon isoprenoid units.

(b)

Plastoquinone

H 3 C

h H 1

h 3 c

(CH 2 -C = C-CH 2 ) 6 _ 10 H

Figure 7.34 ►

Three oxidation states of ubiquinone.

Ubiquinone is reduced in two one-electron
steps via a semiquinone free-radical inter-
mediate. The reactive center of ubiquinone
is shown in red.

Ubiquinone (Q)

CH 3

H I

_c = c-ch 2 ) 6 _ 10

+ e

- pO

Semiquinone anion (*Q 0 )

CH,

(ch 2 — c = c — ch 2 ) 6 _ 10 h

+ 2H 0

+ e 0

- 2 H 0

Ubiquinol (QH 2 )

cn 3

-C = C-CH 2 ) 6 _ 10 H

7.17 Cytochromes 221

Unlike FAD or FMN, ubiquinone and its derivatives cannot accept or donate a pair
of electrons in a single step.

7.16 Protein Coenzymes

Some proteins act as coenzymes. They do not catalyze reactions by themselves but are
required by certain other enzymes. These coenzymes are called either group transfer
proteins or protein coenzymes. They contain a functional group either as part of their
protein backbone or as a prosthetic group. Protein coenzymes are generally smaller
and more heat-stable than most enzymes. They are called coenzymes because they par-
ticipate in many different reactions and associate with a variety of different enzymes.

Some protein coenzymes participate in group transfer reactions or in oxidation-
reduction reactions in which the transferred group is hydrogen or an electron. Metal
ions, iron-sulfur clusters, and heme groups are reactive centers commonly found in
these protein coenzymes. (Cytochromes are an important class of protein coenzymes
that contain heme prosthetic groups. See Section 7.17.) Several protein coenzymes have
two reactive thiol side chains that cycle between their dithiol and disulfide forms. For
example, thioredoxins have cysteines three residues apart ( — Cys — X — X — Cys — ). The
thiol side chains of these cysteine residues undergo reversible oxidation to form the
disulfide bond of a cystine unit. We will encounter thioredoxins as reducing agents
when we examine the citric acid cycle (Chapter 13), photosynthesis (Chapter 15), and
deoxyribonucleotide synthesis (Chapter 18). The disulfide reactive center of thiore-
doxin is on the surface of the protein where it is accessible to the active sites of appro-
priate enzymes (Figure 7.35 ).

Ferredoxin is another common oxidation-reduction coenzyme. It contains two
iron-sulfur clusters that can accept or donate electrons (Figure 7.36 ).

Some other protein coenzymes contain firmly bound coenzymes or portions of
coenzymes. In Escherichia coli , a carboxyl carrier protein containing covalently bound
biotin is one of three protein components of acetyl CoA carboxylase that catalyzes the
first committed step of fatty acid synthesis. (In animal acetyl CoA carboxylases, the
three protein components are fused into one protein chain.) ACP, introduced in Section 7.6,
contains a phosphopantetheine moiety as its reactive center. The reactions of ACP
therefore resemble those of coenzyme A. ACP is a component of all fatty acid synthases
that have been tested. A protein coenzyme necessary for the degradation of glycine in
mammals, plants, and bacteria (Chapter 17) contains a molecule of covalently bound
lipoamide as a prosthetic group.

7.17 Cytochromes

Cytochromes are heme-containing protein coenzymes whose Fe(III) atoms undergo
reversible one-electron reduction. Some structures of cytochromes were shown
in Figures 4.21 and 4.24b. Cytochromes are classified as a, b , and c on the basis of
their visible absorption spectra. The absorption spectra of reduced and oxidized
cytochrome c are shown in Figure 7.37. Although the most strongly absorbing band is
the Soret (or y) band, the band labeled a is used to characterize cytochromes as either
a, b , or c. Cytochromes in the same class may have slightly different spectra; therefore,
a subscript number denoting the peak wavelength of the a absorption band of the
reduced cytochrome often differentiates the cytochromes of a given class (e.g.,
cytochrome fr 56 o). Wavelengths of maximum absorption for reduced cytochromes are
given in Table 7.3.

Figure 7.37 ►

Comparison of the absorption spectra of oxidized (red) and reduced (blue) horse cytochrome c. The re-
duced cytochrome has three absorbance peaks, designated a, ft, and y On oxidation, the Soret (or y)
band decreases in intensity and shifts to a slightly shorter wavelength, whereas the a and p peaks
disappear, leaving a single broad band of absorbance.

The strength of coenzyme oxidizing
agents (standard reduction potential)
is described in Section 10.9.

▲ Figure 7.35

Oxidized thioredoxin. Note that the cystine
group is on the exposed surface of the pro-
tein. The sulfur atoms are shown in yellow.
See Figure 4.24m for another view of thiore-
doxin. [PDB 1ERU].

▲ Figure 7.36

Ferredoxin. This ferredoxin from Pseudomonas
aeruginosa contains two [4 Fe-4 S] iron-
sulfur clusters that can be oxidized and re-
duced. Ferredoxin is a common cosubstrate
in many oxidation-reduction reactions.

[PDB 2FG0]

150-

Soret band (or y)

220 300 400 500 600

Wavelength (nm)

222

CHAPTER 7 Coenzymes and Vitamins

Table 7.3 Absorption maxima (in nm) of major spectral bands in the visible
absorption spectra of the reduced cytochromes

Absorption band

Heme protein

Cytochrome c

550-558

521-527

415-423

Cytochrome b

555-567

526-546

408-449

Cytochrome a

592-604

Absent

439-443

The classes have slightly different heme prosthetic groups (Figure 7.38 ). The heme
of fr-type cytochromes is the same as that of hemoglobin and myoglobin (Figure 4.44).
The heme of cytochrome a has a 17-carbon hydrophobic chain at C-2 of the porphyrin
ring and a formyl group at C-8, whereas the fr-type heme has a vinyl group attached to
C-2 and a methyl group at C-8. In c-type cytochromes, the heme is covalently attached
to the apoprotein by two thioether linkages formed by addition of the thiol groups of
two cysteine residues to vinyl groups of the heme.

The tendency to transfer an electron to another substance, measured as a reduction
potential, varies among individual cytochromes. The differences arise from the different
environment each apoprotein provides for its heme prosthetic group. The reduction
potentials of iron-sulfur clusters also vary widely depending on the chemical and physi-
cal environment provided by the apoprotein. The range of reduction potentials among
prosthetic groups is an important feature of membrane- associated electron transport
pathways (Chapter 14) and photosynthesis (Chapter 15).

CH 3

Figure 7.38 ► l_l |

Heme groups of (a) cytochrome a, (a) CH 2 — (CH 2 — C = C — CH 2 ) 3 — H

(b) cytochrome b, and (c) cytochrome c.

Summary 223

BOX 7.5 NOBEL PRIZES FOR VITAMINS AND COENZYMES

The discovery of vitamins in the first part of the 20th century
stimulated an enormous amount of biochemistry research.
What were these mysterious chemicals that seemed essential
for life? Why were they essential?

We now take vitamins and coenzymes for granted but
that doesn’t do justice to the workers who discovered their
role in metabolism. Here’s a list of the scientists who received
Nobel Prizes for their work on vitamins and coenzymes.

Chemistry 1928: Adolf Otto Reinhold Windaus “for the serv-
ices rendered through his research into the constitution of the
sterols and their connection with the vitamins.”

Physiology or Medicine 1929: Christiaan Eijkman “for his
discovery of the antineuritic vitamin.” Sir Frederick Gow-
land Hopkins “for his discovery of the growth-stimulating
vitamins.”

Chemistry 1937: Paul Karrer “for his investigations on
carotenoids, flavins and vitamins A and B 2 .” Walter Norman
Haworth “for his investigations on carbohydrates and vita-
min C.”

Physiology or Medicine 1937: Albert von Szent-Gyorgyi
Nagyrapolt “for his discoveries in connection with the bio-
logical combustion processes, with special reference to vita-
min C and the catalysis of fumaric acid.”

Chemistry 1938: Richard Kuhn “for his work on carotenoids
and vitamins.”

Physiology or Medicine 1943: Henrik Carl Peter Dam “for
his discovery of vitamin K.” Edward Adelbert Doisy “for his
discovery of the chemical nature of vitamin K.”

Physiology or Medicine 1953: Fritz Albert Lipmann “for his
discovery of co-enzyme A and its importance for intermedi-
ary metabolism.”

Chemistry 1964: Dorothy Crowfoot Hodgkin “for her deter-
minations by X-ray techniques of the structures of important
biochemical substances.”

Chemistry 1970: Luis F. Leloir “for his discovery of sugar nu-
cleotides and their role in the biosynthesis of carbohydrates.”

Chemistry 1997: Paul D. Boyer and John E. Walker “for their
elucidation of the enzymatic mechanism underlying the syn-
thesis of adenosine triphosphate (ATP).”

▲ Nobel Medals. Chemistry (left), Physiology or Medicine (right).

Summary

1. Many enzyme- catalyzed reactions require cofactors. Cofactors in-
clude essential inorganic ions and group-transfer reagents called
coenzymes. Coenzymes can either function as cosubstrates or re-
main bound to enzymes as prosthetic groups.

2. Inorganic ions, such as K®, Mg®, Ca®, Zn®, and Fe®, may
participate in substrate binding or in catalysis.

3. Some coenzymes are synthesized from common metabolites; oth-
ers are derived from vitamins. Vitamins are organic compounds
that must be supplied in small amounts in the diets of humans
and other animals.

4. The pyridine nucleotides, NAD© and NADP©, are coenzymes
for dehydrogenases. Transfer of a hydride ion (H®) from a spe-
cific substrate reduces NAD© or NADP© to NADH or NADPH,
respectively, and releases a proton.

5. The coenzyme forms of riboflavin — FAD and FMN — are tightly
bound as prosthetic groups. FAD and FMN are reduced by
hydride (two-electron) transfers to form FADH 2 and FMNH 2 , re-
spectively. The reduced flavin coenzymes donate electrons one or
two at a time.

6. Coenzyme A, a derivative of pantothenate, participates in acyl-
group-transfer reactions. Acyl carrier protein is required in the
synthesis of fatty acids.

7. The coenzyme form of thiamine is thiamine diphosphate (TDP),
whose thiazolium ring binds the aldehyde generated on decar-
boxylation of an a-keto acid substrate.

8. Pyridoxal 5 '-phosphate is a prosthetic group for many enzymes
in amino acid metabolism. The aldehyde group at C-4 of PLP
forms a Schiff base with an amino acid substrate, through which
it stabilizes a carbanion intermediate.

9. Vitamin C is a vitamin but not a coenzyme. It’s a substrate in
several reactions including those required in the synthesis of
collagen. Vitamin C deficiency causes scurvy. Primates need an
external source of vitamin C because they have lost one of the
key enzymes required for its synthesis. The gene for this enzyme
is a pseudogene in certain primate genomes.

10. Biotin, a prosthetic group for several carboxylases and carboxyl-
transferases, is covalently linked to a lysine residue at the enzyme
active site.

11. Tetrahydrofolate is a reduced derivative of folate and participates
in the transfer of one-carbon units at the oxidation levels of
methanol, formaldehyde, and formic acid. Tetrahydrobiopterin is
a reducing agent in some hydroxylation reactions.

12. The coenzyme forms of cobalamin — adenosylcobalamin and
methylcobalamin — contain cobalt and a corrin ring system.
These coenzymes participate in a few intramolecular rearrange-
ments and methylation reactions.

13. Lipoamide, a prosthetic group for a-keto acid dehydrogenase
multienzyme complexes, accepts an acyl group, forming a thioester.

14. The four fat-soluble, or lipid, vitamins are A, D, E, and K. These
vitamins have diverse functions.

224 CHAPTER 7 Coenzymes and Vitamins

15. Ubiquinone is a lipid- soluble electron carrier that transfers elec-
trons one or two at a time.

16. Some proteins, such as acyl carrier protein and thioredoxin, act as
coenzymes in group-transfer reactions or in oxidation-reduction
reactions in which the transferred group is hydrogen or an electron.

Problems

1. For each of the following enzyme-catalyzed reactions, determine
the type of reaction and the coenzyme that is likely to participate.

OH O

(a) CH 3 — CH— COO© » CH 3 — C— COO©

17. Cytochromes are small, heme- containing protein coenzymes that
participate in electron transport. They are differentiated by their
absorption spectra.

O O

ii n ii

(b) ch 3 — ch 2 — c— coo© » ch 3 — ch 2 — C — H + co 2

o o

11 n n 11

CH 3 O O

(d) ©OOC— CH — C— S-CoA > ©OOC— CH 2 — CH 2 — C —S-CoA

(e) CH 3 — CH— TPP + HS-CoA » CH 3 — C — S-CoA + TPP

2. List the coenzymes that

(a) participate as oxidation-reduction reagents.

(b) act as acyl carriers.

(d) transfer groups to and from amino acids.

(e) are involved in carboxylation or decarboxylation reactions.

3. In the oxidation of lactate to pyruvate by lactate dehydrogenase
(LDH), NAD® is reduced in a two-electron transfer process from
lactate. Since two protons are removed from lactate as well, is it cor-
rect to write the reduced form of the coenzyme as NADH 2 ? Explain.

h 3 c— c— coo©

L-Lactate

Pyruvate

4. Succinate dehydrogenase requires FAD to catalyze the oxidation
of succinate to fumarate in the citric acid cycle. Draw the isoalloxazine
ring system of the cofactor resulting from the oxidation of succi-
nate to fumarate and indicate which hydrogens in FADH 2 are
lacking in FAD.

©ooc— ch 2 — ch 2 — coo©

Succinate

Fumarate

©OOC — CH = CH — COO©

5. What is the common structural feature of NAD®, FAD, and
coenzyme A?

6. Certain nucleophiles can add to C-4 of the nicotinamide ring of
NAD®, in a manner similar to the addition of a hydride in the re-
duction of NAD® to NADH. Isoniazid is the most widely used
drug for the treatment of tuberculosis. X-ray studies have shown
that isoniazid inhibits a crucial enzyme in the tuberculosis bac-
terium where a covalent adduct is formed between the carbonyl
of isoniazid and the 4' position of the nicotinamide ring of a
bound NAD® molecule. Draw the structure of this NAD-isoni-
azid inhibitory adduct.

Isoniazid

NHNH 2

7. A vitamin B 6 deficiency in humans can result in irritability,
nervousness, depression, and sometimes convulsions. These
symptoms may result from decreased levels of the neurotrans-
mitters serotonin and norepinephrine, which are metabolic de-
rivatives of tryptophan and tyrosine, respectively. How could a
deficiency of vitamin B 6 result in decreased levels of serotonin
and norepinephrine?

Problems 225

Serotonin

Norepinephrine

8. Macrocytic anemia is a disease in which red blood cells mature
slowly due to a decreased rate of DNA synthesis. The red blood cells
are abnormally large (macrocytic) and are more easily ruptured.
How could the anemia be caused by a deficiency of folic acid?

9 . A patient suffering from methylmalonic aciduria (high levels of
methylmalonic acid) has high levels of homocysteine and low
levels of methionine in the blood and tissues. Folic acid levels are
normal.

(a) What vitamin is likely to be deficient?

(b) How could the deficiency produce the symptoms listed above?

10 . Alcohol dehydrogenase (ADH) from yeast is a metalloenzyme
that catalyzes the NAD® -dependent oxidation of ethanol to ac-
etaldehyde. The mechanism of yeast ADH is similar to that of
lactate dehydrogenase (LDH) (Figure 7.9) except that the zinc
ion of ADH occupies the place of His- 195 in LDH.

(a) Draw a mechanism for the oxidation of ethanol to acetalde-
hyde by yeast ADH.

(b) Does ADH require a residue analogous to Arg-171 in LDH?

11. In biotin- dependent transcarboxylase reactions, an enzyme trans-
fers a carboxyl group between substrates in a two-step process
without the need for ATP or bicarbonate. The reaction catalyzed
by the enzyme methylmalonyl CoA- pyruvate transcarboxylase is
shown below. Draw the structures of the products expected from
the first step of the reaction.

ch 3 o o

o 1 11 11

©OOC— CH— C— S-CoA + CH 3 — C— COO©

Methylmalonyl CoA Pyruvate

CH 3 — CH 2 — C— S-CoA
Propionyl CoA

©OOC— ch 2 — c — coo©
Oxaloacetate

12 . (a) Histamine is produced from histidine by the action of a de-
carboxylase. Draw the external aldimine produced by the re-
action of histidine and pyridoxal phosphate at the active site
of histidine decarboxylase.

(b) Since racemization of amino acids by PLP-dependent en-
zymes proceeds via Schiff base formation, would racemiza-
tion of L-histidine to D-histidine occur during the histidine
decarboxylase reaction?

13 . (a) Thiamine pyrophosphate is a coenzyme for oxidative decar-
boxylation reactions in which the keto carbonyl carbon is ox-
idized to an acid or an acid derivative. Oxidation occurs by
removal of two electrons from a resonance- stabilized carban-
ion intermediate. What is the mechanism for the reaction
pyruvate + HS-CoA —> acetyl CoA + C0 2 , beginning from
the resonance-stabilized carbanion intermediate formed after
decarboxylation (Figure 7.15) (such as a thioester in the case
below)?

(b) Pyruvate dehydrogenase (PDH) is an enzyme complex that
catalyzes the oxidative decarboxylation of pyruvate to acetyl
CoA and C0 2 in a multistep reaction. The oxidation and
acetyl-group transfer steps require TDP and lipoic acid in
addition to other coenzymes. Draw the chemical structures
for the molecules in the following two steps in the PDH
reaction.

HETDP + lipoamide » acetyl-TDP + dihydrolipoamide »

TDP + acetyl-dihydrolipoamide

(c) In a transketolase enzyme TDP-dependent reaction, the
resonance-stabilized carbanion intermediate shown adjacent
is generated as an intermediate. This intermediate is then in-
volved in a condensation reaction (resulting in C — C bond
formation) with the aldehyde group of erythrose 4-phos-
phate (E4P) to form fructose 6-phosphate (F6P). Starting
from the carbanion intermediate, show a mechanism for this
transketolase reaction. (Fischer projections of carbohydrate
structures are sometimes drawn as shown here.)

H— C— OH

TDP

H— C— OH

hoch 2 — c — oh

ch 2 opo 3 ©

Intermediate Erythrose

4-phosphate

CH 2 OH
C = 0

HO— C — H

H— C— OH

CH 2 0P0 3 ©

Fructose

6-phosphate

226 CHAPTER 7 Coenzymes and Vitamins

Selected Readings

Metal Ions

Berg, J. M. (1987). Metal ions in proteins: struc-
tural and functional roles. Cold Spring Harbor
Symp. Quant. Biol 52:579-585.

Rees, D. C. (2002). Great metalloclusters in enzy-
mology. Annu. Rev. Biochem. 71: 221-246.

Specific Cofactors

Banerjee, R., and Ragsdale, S.W. (2003). The many
faces of vitamin B 12 : catalysis by cobalmin-
dependent enzymes. Annu. Rev. Biochem.
72:209-247.

Bellamacina, C. R. (1996). The nicotinamide
dinucleotide binding motif: a comparison of
nucleotide binding proteins. FASEB J.
10:1257-1268.

Blakley, R. L., and Benkovic, S. J., eds. (1985).
Folates and Pterins, Vol. 1 andVol. 2. (New York:
John Wiley 8c Sons).

Chiang, R K., Gordon, R. K., Tal, J., Zeng, G. C.,
Doctor, B. P., Pardhasaradhi, K., and McCann,

P. P. (1996). S-Adenosylmethionine and methylation.
FASEB J. 10:471-480.

Coleman, J. E. (1992). Zinc proteins: enzymes, stor-
age proteins, transcription factors, and replication
proteins. Annu. Rev. Biochem. 61:897-946.

Ghisla, S., and Massey, V. (1989). Mechanisms of
flavoprotein-catalyzed reactions. Eur. J. Biochem.
181:1-17.

Hayashi, H., Wada, H., Yoshimura, T., Esaki, N.,
and Soda, K. (1990). Recent topics in pyridoxal
5 '-phosphate enzyme studies. Annu. Rev. Biochem.
59:87-110.

Jordan, F. (1999). Interplay of organic and biologi-
cal chemistry in understanding coenzyme mecha-
nisms: example of thiamin diphosphate-dependent
decarboxylations of 2-oxo acids. FEBS Lett.
457:298-301.

Jordan, F., Li, EL, and Brown, A. (1999). Remark-
able stabilization of zwitterionic intermediates
may account for a billion-fold rate acceleration by
thiamin diphosphate- dependent decarboxylases.
Biochem. 38:6369-6373.

Jurgenson, C. T., Begley, T. P. and Ealick, S. E.
(2009). The structural and biochemical founda-
tions of thiamin biosynthesis. Ann. Rev. Biochem.
78:569-603.

Knowles, J. R. (1989). The mechanism of biotin-
dependent enzymes .Annu. Rev. Biochem. 58:195-221.

Ludwig, M. L., and Matthews, R. G. (1997).
Structure-based perspectives on B 12 -dependent
enzymes .Annu. Rev. Biochem. 66:269-313.

Palfey, B. A., Moran, G. R., Entsch, B., Ballou, D. P.,
and Massey, V. (1999). Substrate recognition by
“password” in p-hydroxybenzoate hydroxylase.
Biochem. 38:1153-1158.

NAD-Binding Motifs

Bellamacina, C. R. (1996). The nictotinamide
d inucleotide binding motif: a comparison of nu-
cleotide binding proteins. FASEB /. 10:1257-1269.

Rossman, M. G., Liljas, A., Branden, C.-L, and
Banaszak, L. J. (1975). Evolutionary and structural
relationships among dehydrogenases. In The Enzymes.
Vol. 11, Part A, 3rd ed., P. D., Boyer, ed. (New York:
Academic Press), pp. 61-102.

Wilks, H. M., Hart, K. W., Feeney, R., Dunn, C. R.,
Muirhead, H., Chia, W. N., Barstow, D. A., Atkin-
son, T., Clarke, A. R., and Holbrook, J. J. (1988).

A specific, highly active malate dehydrogenase by
redesign of a lactate dehydrogenase framework.
Science 242:1541-1544.

o c

_ o

° o o o

° o

o o

° c

o o

Carbohydrates

C arbohydrates (also called saccharides) are — on the basis of mass — the most
abundant class of biological molecules on Earth. Although all organisms can
synthesize carbohydrate, much of it is produced by photosynthetic organ-
isms, including bacteria, algae, and plants. These organisms convert solar energy to
chemical energy that is then used to make carbohydrate from carbon dioxide. Carbo-
hydrates play several crucial roles in living organisms. In animals and plants, carbohy-
drate polymers act as energy storage molecules. Animals can ingest carbohydrates
that can then be oxidized to yield energy for metabolic processes. Polymeric carbohy-
drates are also found in cell walls and in the protective coatings of many organisms.
Other carbohydrate polymers are marker molecules that allow one type of cell to rec-
ognize and interact with another type. Carbohydrate derivatives are found in a num-
ber of biological molecules, including some coenzymes (Chapter 7) and the nucleic
acids (Chapter 19).

The name carbohydrate , “hydrate of carbon,” refers to their empirical formula
(CH 2 0) n , where n is 3 or greater ( n is usually 5 or 6 but can be up to 9). Carbohydrates
can be described by the number of monomeric units they contain. Monosaccharides are
the smallest units of carbohydrate structure. Oligosaccharides are polymers of two to
about 20 monosaccharide residues. The most common oligosaccharides are disaccha-
rides, which consist of two linked monosaccharide residues. Polysaccharides are
polymers that contain many (usually more than 20) monosaccharide residues.
Oligosaccharides and polysaccharides do not have the empirical formula (CH 2 0) n be-
cause water is eliminated during polymer formation. The term glycan is a more general
term for carbohydrate polymers. It can refer to a polymer of identical sugars (homoglycan)
or of different sugars (heteroglycan).

Glycoconjugates are carbohydrate derivatives in which one or more carbohydrate
chains are linked covalently to a peptide, protein, or lipid. These derivatives include pro-
teoglycans, peptidoglycans, glycoproteins, and glycolipids.

In this chapter, we discuss nomenclature, structure, and function of monosaccha-
rides, disaccharides, and the major homoglycans — starch, glycogen, cellulose, and

Molecular biology has dealt largely
on the triad of DNA , RNA and pro-
tein. Biochemistry is concerned with
all the molecules of the cell. Excluded
from the province of molecular biol-
ogy have been most of the structures
and functions essential for growth
and maintenance: carbohydrates ,
coenzymes ; lipids , and membranes.

— Arthur Korn berg
"For the love of enzymes: the
odyssey of a biochemist" (1 989)

Photosynthesis is described in detail in
Chapter 15.

Top: Darkling beetle. The exoskeletons of insects contain chitin, a homoglycan.

227

228 CHAPTER 8 Carbohydrates

KEY CONCEPT

A Fischer projection is a convention
designed to convey information about the
stereochemistry of a molecule. It does not
resemble the actual conformation of the
molecule in solution.

Stereo

view

H — C — OH
C

Fischer

projection

For each chiral carbon atom in a
Fischer projection the vertical bonds
project into the plane of the page and
the horizontal bonds project upward
toward the viewer.

Mirror plane

L-Glyceraldehyde D-Glyceraldehyde
▲ Figure 8.2

View of L-glyceraldehyde (left) and o-glycer-
aldehyde (right). These molecules are drawn
in a conformation that corresponds to the
Fischer projections in Figure 8.1.

chitin. We then consider proteoglycans, peptidoglycans, and glycoproteins, all of which
contain heteroglycan chains.

8.1 Most Monosaccharides Are Chiral Compounds

Monosaccharides are water-soluble, white, crystalline solids that have a sweet taste. Ex-
amples include glucose and fructose. Chemically, monosaccharides are polyhydroxy
aldehydes, or aldoses, or polyhydroxy ketones, or ketoses. They are classified by their
type of carbonyl group and their number of carbon atoms. As a rule, the suffix -ose is
used in naming carbohydrates, although there are a number of exceptions. All mono-
saccharides contain at least three carbon atoms. One of these is the carbonyl carbon,
and each of the remaining carbon atoms bears a hydroxyl group. In aldoses, the most
oxidized carbon atom is designated C-l and is drawn at the top of a Fischer projection.
In ketoses, the most oxidized carbon atom is usually C-2.

We’ve encountered Fischer projections before but now it’s time to present the con-
vention in more detail. A Fischer projection is a two-dimensional representation of a
three-dimensional molecule. It is designed to preserve information about the stereo-
chemistry of a molecule. In a Fischer projection of sugars, the C-l atom is always at
the top of the figure. For each separate chiral carbon atom, the two horizontal bonds
project upward from the page toward you. The two vertical bonds project downward
into the page. Remember, this applies to each chiral carbon atom, so in a carbohydrate
with multiple carbon atoms the Fischer projection represents a molecule that curls back
into the page. For longer molecules, the top and bottom groups may even come in vir-
tual contact, forming a loop. The Fischer projection is a convention for preserving
stereochemical information; it does not represent a realistic model of how a molecule
might look in solution.

The smallest monosaccharides are trioses, or three-carbon sugars. One- or two-carbon
compounds having the general formula (CH 2 0)„ do not have properties typical of car-
bohydrates (such as sweet taste and the ability to crystallize). The aldehydic triose, or
aldotriose, is glyceraldehyde (Figure 8.1a). Glyceraldehyde is chiral because its central
carbon, C-2, has four different groups attached to it, (Section 3.1). The ketonic triose, or
ketotriose, is dihydroxyacetone (Figure 8.1b). It is achiral because it has no asymmetric
carbon atom. All other monosaccharides, longer- chain versions of these two sugars, are
chiral.

The stereoisomers d- and L-glyceraldehyde are shown as ball-and-stick models in
Figure 8.2. Chiral molecules are optically active; that is, they rotate the plane of polar-
ized light. The convention for designating D and L isomers was originally based on the
optical properties of glyceraldehyde. The form of glyceraldehyde that caused rotation to
the right (dextrorotatory) was designated d and the form that caused rotation to the left
(levorotatory) was designated l. Structural knowledge was limited when this conven-
tion was established in the late 19th century so the configurations for the enantiomers
of glyceraldehyde were assigned arbitrarily, with a 50% probability of error. X-ray
crystallographic experiments later proved that the original structural assignments were
correct.

H /O

\ f

H /O

\ S

(b)

CH 2 OH

HO — C — H

H — C — OH

c=o

CH 2 OH

ch 2 oh

L-Glyceraldehyde

D-Glyceraldehyde

Dihydroxyacetone

▲ Figure 8.1

Fischer projections of (a) glyceraldehyde and (b) dihydroxyacetone. The designations l (for left) and d
(for right) for glyceraldehyde refer to the configuration of the hydroxyl group of the chiral carbon
(C-2). Dihydroxyacetone is achiral.

8.1 Most Monosaccharides Are Chiral Compounds 229

Aldotriose

H — C — OH

3 CH 2 OH

D-Glyceraldehyde

H — C — OH

4 ch 2 oh

D-Erythrose

Aldotetroses

H s

HO— C — H

H — C — OH

ch 2 oh

D-Threose

Aldopentoses

H ",c

H ^°

H “l

— OH

HO — c — H

H — C — OH

HO — C — H

H-.

— OH

H — C — OH

HO — C — H

H — C

4 i

— OH

H — C — OH

5 ch 2 oh

CH 2 OH

D-Ribose

D-Arabinose

D-Xylose

D-Lyxose

i i i i

Aldohexoses

i i

H ^°

— u-
1

HO— C — H

1 1

h— c— oh ho— c — h

i i

H — C— OH HO— C — H

H — C— OH HO— C— H

H — C— OH

3 |

H — C— OH

HO— c — H HO— c— H

H — C— OH H — C— OH

HO— C — H

H — C— OH

4 |

H — C— OH

H — C— OH H — C— OH

HO— C— H HO— C— H

HO— C— H

H — C— OH

5 |

H — C— OH

H — C— OH H — C— OH

H — C— OH

ch 2 oh

ch 2 oh ch 2 oh

CH 2 OH

D-Allose

D-Altrose

D-Glucose D-Mannose

D-Gulose D-ldose

D-Galactose D-Talose

▲ Figure 8.3

Fischer projections of the three- to six-carbon D-aldoses. The aldoses shown in blue are the most important in our study of biochemistry.

Longer aldoses and ketoses can be regarded as extensions of glyceraldehyde and di-
hydroxyacetone, respectively, with chiral H — C — OH groups inserted between the car-
bonyl carbon and the primary alcohol group. Figure 8.3 shows the complete list of the
names and structures of the tetroses (four-carbon aldoses), pentoses (five-carbon al-
doses), and hexoses (six-carbon aldoses) related to D- glyceraldehyde. Many of these
monosaccharides are not synthesized by most organisms and we will not encounter
them again in this book.

Note that the carbon atoms are numbered from the carbon of the aldehyde group
that is assigned the number 1. By convention, sugars are said to have the D configuration
when the configuration of the chiral carbon with the highest number — the chiral carbon
most distant from the carbonyl carbon — is the same as that of C-2 of D- glyceraldehyde

230 CHAPTER 8 Carbohydrates

Figure 8.4 ►

l- and o-glucose. Fischer projections (left)
showing that l- and D-glucose are mirror
images. Conformation of the extended form
of D-glucose in solution.

(i.e., the — OH group attached to this carbon atom is on the right side in a Fischer pro-
jection). The arrangement of asymmetric carbon atoms is unique for each monosac-
charide, giving each its distinctive properties. Except for glyceraldehyde (which was
used as the standard), there is no predictable association between the absolute configu-
ration of a sugar and whether it is dextrorotatory or levorotatory.

It is mostly the D enantiomers that are synthesized in living cells — just as the
L enantiomers of amino acids are more common. The L enantiomers of the 15 aldoses in
Figure 8.3 are not shown. Recall that pairs of enantiomers are mirror images; in other
words, the configuration at each chiral carbon is opposite. For example, the hydroxyl
groups bound to carbon atoms 2, 3, 4, and 5 of D-glucose point right, left, right, and
right, respectively, in the Fischer projection; those of L- glucose point left, right, left, and
left (Figure 8.4).

The three-carbon aldose, glyceraldehyde, has only a single chiral atom (C-2) and
therefore only two stereoisomers. There are four stereoisomers for aldotetroses (d- and
L-erythrose and D- and L-threose) because erythrose and threose each possess two chiral
carbon atoms. In general, there are 2 n possible stereoisomers for a compound with n
chiral carbons. Aldohexoses, which possess four chiral carbons, have a total of 2 4 , or 16,
stereoisomers (the eight D aldohexoses in Figure 8.3 and their L enantiomers).

Sugar molecules that differ in configuration at only one of several chiral centers are
called epimers. For example, D-mannose and D-galactose are epimers of D-glucose (at C-2
and C-4, respectively), although they are not epimers of each other (Figure 8.3).

Longer- chain ketoses (Figure 8.5) are related to dihydroxyacetone in the same way
that longer-chain aldoses are related to glyceraldehyde. Note that a ketose has one fewer
chiral carbon atom than the aldose of the same empirical formula. For example, there
are only two stereoisomers for the one ketotetrose (d- and L-erythrulose), and four
stereoisomers for ketopentoses (d- and L-xylulose and d- and L-ribulose). Ketotetrose
and ketopentoses are named by inserting -ul- in the name of the corresponding aldose.
For example, the ketose xylulose corresponds to the aldose xylose. This nomenclature
does not apply to the ketohexoses (tagatose, sorbose, psicose, and fructose) because they
have traditional (trivial) names.

Mirror

plane

H /O

H ,0

\ f

\ s

-u-

H— 2 C— OH

- u-
1

- u-

- u-
1

H— 4 C— OH

-u-

CH 2 OH

6 ch 2 oh

L-Glucose

D-Glucose

°M

«*»

D-Glucose

8.2 Cyclization of Aldoses and Ketoses

The optical behavior of some monosaccharides suggests they have one more chiral
carbon atom than is evident from the structures shown in Figures 8.3 and 8.5.
D-Glucose, for example, exists in two forms that contain five (not four) asymmetric carbons.
The source of this additional asymmetry is an intramolecular cyclization reaction that
produces a new chiral center at the carbon atom of the carbonyl group. This cyclization
resembles the reaction of an alcohol with an aldehyde to form a hemiacetal or with a
ketone to form a hemiketal (Figure 8.7).

The carbonyl carbon of an aldose containing at least five carbon atoms or of a ke-
tose containing at least six carbon atoms can react with an intramolecular hydroxyl

8.2 Cyclization of Aldoses and Ketoses 231

Ketotriose

CH 2 OH
C =0

ch 2 oh

Dihydroxyacetone

Ketotetrose CH 2 OH

C = 0

H — C — OH

ch 2 oh

D-Erythrulose

Ketopentoses

CH 2 OH

c=o

H — C— OH

ch 2 oh

D-Ribulose

CH 2 OH

C=0

HO — C — H

CH 2 OH

D-Xylulose

Ketohexoses

}

* *
z*

£ *

▲ Who am I? The structures of the d sugars
are shown in Figures 8.3 and 8.5. You can
deduce the structures of the l configurations.
Knowing the convention for Fischer projec-
tions, you should have no trouble identifying
these molecules.

ch 2 oh

c=o

H — C— OH

ch 2 oh

D-Psicose

CH 2 OH

C=0

HO — C — H

H — C— OH

ch 2 oh

D-Fructose

CH 2 OH

C=0

HO — C — H

H — C— OH

ch 2 oh

D-Tagatose

CH 2 OH

C=0

HO — C — H

H — C— OH

ch 2 oh

D-Sorbose

◄ Figure 8.5

Fischer projections of the three- to six-carbon
o-ketoses. The ketoses shown in blue are the
most important in our study of biochemistry.

group to form a cyclic hemiacetal or cyclic hemiketal, respectively. The oxygen atom
from the reacting hydroxyl group becomes a member of the five- or six-membered ring
structures (Figure 8.8).

Because it resembles the six-membered heterocyclic compound pyran (Figure 8.6a),
the six-membered ring of a monosaccharide is called a pyranose. Similarly, because the
five-membered ring of a monosaccharide resembles furan (Figure 8.6b), it is called a
furanose. Note that, unlike pyran and furan, the rings of carbohydrates do not contain
double bonds.

The most oxidized carbon of a cyclized monosaccharide, the one attached to two
oxygen atoms, is referred to as the anomeric carbon. In ring structures, the anomeric car-
bon is chiral. Thus, the cyclized aldose or ketose can adopt either of two configurations
(designated a or /J), as illustrated for D-glucose in Figure 8.8. The a and (3 isomers are
called anomers.

In solution, aldoses and ketoses that form ring structures equilibrate among their vari-
ous cyclic and open-chain forms. At 31°C, for example, D-glucose exists in an equilibrium

▲ Figure 8.6

(a) Pyran and (b) furan.

232

CHAPTER 8 Carbohydrates

(a) |_j©

y O Aldehyde

^11

Alcohol

R— C — H

H ©

Hemiacetal

(chiral)

(b)

Alcohol

Hemiketal

(chiral)

▲ Figure 8.7

Hemiacetal and hemiketal. (a) Reaction of an
alcohol with an aldehyde to form a hemi-
acetal. (b) Reaction of an alcohol with a
ketone to form a hemiketal. The asterisks
indicate the newly formed chiral centers.

mixture of approximately 64% /J- D - glucopyr anose and 36% a-D-glucopyranose, with very
small amounts of the furanose (Figure 8.9 ) and open-chain (Figure 8.4) forms. Similarly,
D-ribose exists as a mixture of approximately 58.5% /3-D-ribopyranose, 21.5% a-D-ribopy-
ranose, 13.5% /3-D-ribofuranose, and 6.5% a-D-ribofuranose, with a tiny fraction in the
open-chain form (Figure 8.10). The relative abundance of the various forms of monosac-
charides at equilibrium reflects the relative stabilities of each form. Although unsubstituted
D-ribose is most stable as the /3- pyranose, its structure in nucleotides (Section 8.5c) is the
( 3 - furanose form.

The ring drawings shown in these figures are called Haworth projections, after
Norman Haworth who worked on the cyclization reactions of carbohydrates and first

Figure 8.8 ►

Cyclization of o-glucose to form glucopyranose.

The Fischer projection (top left) is rearranged
into a three-dimensional representation
(top right). Rotation of the bond between C-4
and C-5 brings the C-5 hydroxyl group close
to the C-l aldehyde group. Reaction of the
C-5 hydroxyl group with one side of C-l gives
a-D-glucopyranose; reaction of the hydroxyl
group with the other side gives /kD-glucopy-
ranose. The glucopyranose products are shown
as Haworth projections in which the lower
edges of the ring (thick lines) project in front
of the plane of the paper and the upper
edges project behind the plane of the paper.
In the a-D-anomer of glucose, the hydroxyl
group at C-l points down; in the /TD-anomer,
it points up.

D-Glucose

(Fischer projection)

H OH

u-D-Glucopyranose
(Haworth projection)

/3-D-Glucopyranose
(Haworth projection)

8.2 Cyclization of Aldoses and Ketoses 233

proposed these representations. He received the Nobel Prize in Chemistry in 1937 for
his work on carbohydrate structure and the synthesis of vitamin C.

A Haworth projection adequately indicates stereochemistry and can be easily re-
lated to a Fischer projection: groups on the right in a Fischer projection point downwards
in a Haworth projection. Because rotation around carbon-carbon bonds is constrained
in the ring structure, the Haworth projection is a much more faithful representation of
the actual conformation of sugars.

By convention, a cyclic monosaccharide is drawn so the anomeric carbon is on the
right and the other carbons are numbered in a clockwise direction. In a Haworth pro-
jection, the configuration of the anomeric carbon atom is designated a if its hydroxyl
group is cis to (on the same side of the ring as) the oxygen atom of the highest-numbered
chiral carbon atom. It is /3 if its hydroxyl group is trans to (on the opposite side
of the ring from) the oxygen attached to the highest-numbered chiral carbon. With
a-D-glucopyranose, the hydroxyl group at the anomeric carbon points down; with
/3-D-glucopyranose, it points up.

Monosaccharides are often drawn in either the a- or /3-D-furanose or the a- or
/3-D-pyranose form. However, you should remember that the anomeric forms of five-
and six- carbon sugars are in rapid equilibrium. Throughout this chapter and the rest
of the book, we draw sugars in the correct anomeric form if it is known. We refer to
sugars in a nonspecific way (e.g., glucose) when we are discussing an equilibrium

▲ Figure 8.9

a-o-glucofuranose (top) and /3-o-glucofuranose
(bottom).

5 CH 2 OH

◄ Figure 8.10

Cyclization of o-ribose to form a- and /3-d-
ribopyranose and a- and /3-o-ribofuranose.

D-Ribose

(Fischer projection)

a-D-Ribopyranose /3-D-Ribopyranose

(Haworth projection) (Haworth projection)

a-D-Ribofuranose /3-D-Ribofuranose

(Haworth projection) (Haworth projection)

234 CHAPTER 8 Carbohydrates

▲ Galactose mutarotase. Mutarotases are en-
zymes that catalyze the interconversion of a
and /3 configurations. This interconversion
involves the breaking and remaking of cova-
lent bonds, which is why they are different
configurations. The enzyme shown here is
galactose mutarotase from Lactococcus
lactis with a molecule of a-D-galactose
in the acitve site. The bottom figure shows
the conformation of this molecule. Can you
identify this conformation? [PDB 1L7K]

mixture of the various anomeric forms as well as the open-chain forms. When we
are discussing a specific form of a sugar, however, we will refer to it precisely (e.g.,
/3-D-gluco pyranose). Also, since the d enantiomers of carbohydrates predominate in
nature, we always assume that a carbohydrate has the D configuration unless specified
otherwise.

8.3 Conformations of Monosaccharides

Haworth projections are commonly used in biochemistry because they accurately
depict the configuration of the atoms and groups at each carbon atom of the sugar’s
backbone. However, the geometry of the carbon atoms of a monosaccharide ring is
tetrahedral (bond angles near 110°), so monosaccharide rings are not actually planar.
Cyclic monosaccharides can exist in a variety of conformations (three-dimensional
shapes having the same configuration). Furanose rings adopt envelope conformations
in which one of the five ring atoms (either C-2 or C-3) is out-of-plane and the remaining
four are approximately coplanar (Figure 8.11). Furanoses can also form twist conformations
where two of the five ring atoms are out-of-plane — one on either side of the plane
formed by the other three atoms. The relative stability of each conformer depends on
the degree of steric interference between the hydroxyl groups. The various conformers
of unsubstituted monosaccharides can rapidly interconvert.

Pyranose rings tend to assume one of two conformations, the chair conformation
or the boat conformation (Figure 8.12). There are two distinct chair conformers and six
distinct boat conformers for each pyranose. The chair conformations minimize steric
repulsion among the ring substituents and are generally more stable than boat confor-
mations. The — H, — OH, and — CH 2 OH substituents of a pyranose ring in the chair
conformation may occupy two different positions. In the axial position the substituent
is above or below the plane of the ring, while in the equatorial position the substituent
lies in the plane of the ring. In pyranoses, five substituents are axial and five are equatorial.
Whether a group is axial or equatorial depends on which carbon atom (C-l or C-4) ex-
tends above the plane of the ring when the ring is in the chair conformation. Figure 8.13
shows the two different chair conformers of /3-D-glucopyranose. The more stable
conformation is the one in which the bulkiest ring substituents are equatorial (top
structure). In fact, this conformation of /3-D-glucose has the least steric strain of any aldo-
hexose. Pyranose rings are occasionally forced to adopt slightly different conformations,
such as the unstable half- chair adopted by a polysaccharide residue in the active site of
lysozyme (Section 6.6).

KEY CONCEPT

Different configurations can only be
formed by breaking and reforming
covalent bonds. Molecules can adopt
different conformations without breaking
covalent bonds.

Figure 8.1 1 ►

Conformations of /J-o-ribofuranose. (a) Haworth
projection, (b) C 2 -endo envelope conformation,
(c) C 3 -endo envelope conformation, (d) Twist
conformation. In the C 2 -endo conformation,
C-2 lies above the plane defined by C-l, C-3,
C-4, and the ring oxygen. In the C 3 -endo
conformation, C-3 lies above the plane de-
fined by C-l, C-2, C-4, and the ring oxygen.
In the twist conformation shown, C-3 lies
above and C-2 lies below the plane defined
by C-l, C-4, and the ring oxygen. The planes
are shown in yellow.

(a)

(c)

Haworth projection

(b)

C 2 -endo envelope conformation
(d)

5 H

HOCH-, VO

C 3 -endo envelope conformation

Twist conformation

8.4 Derivatives of Monosaccharides 235

(a)

Haworth projection

Chair conformation

Boat conformation

(b)

◄ Figure 8.12

Conformations of /J-o-glucopyranose.

(a) Haworth projection, a chair conformation,
and a boat conformation, (b) Bal l-and-stick
model of a chair (left) and a boat (right)
conformation.

8.4 Derivatives of Monosaccharides

There are many known derivatives of the basic monosaccharides. They include poly-
merized monosaccharides, such as oligosaccharides and polysaccharides, as well as sev-
eral classes of nonpolymerized compounds. In this section, we introduce a few mono-
saccharide derivatives, including sugar phosphates, deoxy and amino sugars, sugar
alcohols, and sugar acids.

Like other polymer-forming biomolecules, monosaccharides and their derivatives
have abbreviations used in describing more complex polysaccharides. The accepted ab-
breviations contain three letters, with suffixes added in some cases. The abbreviations
for some pentoses and hexoses and their major derivatives are listed in Table 8.1. We use
these abbreviations later in this chapter.

A. Sugar Phosphates

Monosaccharides are often converted to phosphate esters. Figure 8.14 shows the struc-
tures of several of the sugar phosphates we will encounter in our study of carbohydrate
metabolism. The triose phosphates, ribose 5-phosphate, and glucose 6-phosphate are
simple alcohol-phosphate esters. Glucose 1 -phosphate is a hemiacetal phosphate, which
is more reactive than an alcohol phosphate. The ability of UDP- glucose to act as a glu-
cosyl donor (Section 7.3) is evidence of this reactivity.

▲ Figure 8.13

The two chair conformers of /?-D-glucopyranose.

The top conformer is more stable.

B. Deoxy Sugars

The structures of two deoxy sugars are shown in Figure 8.15. In these derivatives, a
hydrogen atom replaces one of the hydroxyl groups in the parent monosaccharide.
2-Deoxy-D-ribose is an important building block for DNA. L-Fucose (6-deoxy-L-galac-
tose) is widely distributed in plants, animals, and microorganisms. Despite its unusual
L configuration, fucose is derived metabolically from D-mannose.

C. Amino Sugars

In a number of sugars, an amino group replaces one of the hydroxyl groups in the parent
monosaccharide. Sometimes the amino group is acetylated. Three examples of amino

236 CHAPTER 8 Carbohydrates

Table 8.1 Abbreviations for some monosac-
charides and their derivatives

Monosaccharide

or derivative Abbreviation

Pentoses

Ribose

Rib

Xylose

Xyl

Hexoses

Fructose

Fru

Galactose

Gal

Glucose

Glc

Mannose

Man

Deoxy sugars

Abequose

Abe

Fucose

Fuc

Amino sugars

Glucosamine

GlcN

Galactosamine

GaIN

N-Acetylglucosamine

GIcNAc

N- Acetylgalactosamine
N-Acetylneuraminic acid

GalNAc

NeuNAc

N-Acetyl mu ramie acid

MurNAc

Sugar acids

Glucuronic acid

GlcUA

Iduronic acid

IdoA

ch 2 oh

c = o

1 ©

ch 2 opo 3 ^

■c'°

-c — OH

ch 2 opo

Dihydroxyacetone

phosphate

D-Glyceraldehyde

3-phosphate

▲ Figure 8.14

Structures of several metabolically important sugar phosphates.

sugars are shown in Figure 8.16. Amino sugars formed from glucose and galactose
commonly occur in glycoconjugates. N-Acetylneuraminic acid (NeuNAc) is an acid
formed from N-acetylmannosamine and pyruvate. When this compound cyclizes to
form a pyranose, the carbonyl group at C-2 (from the pyruvate moiety) reacts with the
hydroxyl group of C-6. NeuNAc is an important constituent of many glycoproteins and
of a family of lipids called gangliosides (Section 9.5). Neuraminic acid and its deriva-
tives, including NeuNAc, are collectively known as sialic acids.

/3-2-Deoxy-D-ribose

u-L-Fucose

(6-Deoxy-L-galactose)

▲ Figure 8.15

Structures of the deoxy sugars 2-deoxy-o-ribose
and L-fucose.

D. Sugar Alcohols

In a sugar alcohol, the carbonyl oxygen of the parent monosaccharide has been reduced,
producing a polyhydroxy alcohol. Figure 8.17 shows three examples of sugar alcohols.
Glycerol and rayo-inositol are important components of lipids (Section 10.4). Ribitol is
a component of flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD)
(Section 7.4). In general, sugar alcohols are named by replacing the suffix -ose of the
parent monosaccharides with -itol.

E. Sugar Acids

Sugar acids are carboxylic acids derived from aldoses, either by oxidation of C- 1 (the
aldehydic carbon) to yield an aldonic acid or by oxidation of the highest-numbered
carbon (the carbon bearing the primary alcohol) to yield an alduronic acid. The struc-
tures of the aldonic and alduronic derivatives of glucose — gluconate and glucuronate —
are shown in Figure 8.18. Aldonic acids exist in the open-chain form in alkaline solution
and form lactones (intramolecular esters) on acidification. Alduronic acids can exist as
pyranoses and therefore possess an anomeric carbon. Note that N-acetylneuraminic
acid (Figure 8.16) is a sugar acid as well as an amino sugar. Sugar acids are important
components of many polysaccharides. L-Ascorbic acid or vitamin C, is an enediol of a
lactone derived from D-glucuronate (Section 7.9).

8.5 Disaccharides and Other Glycosides

The glycosidic bond is the primary structural linkage in all polymers of monosaccha-
rides. A glycosidic bond is an acetal linkage in which the anomeric carbon of a sugar is
condensed with an alcohol, an amine, or a thiol. As a simple example, glucopyranose

8.5 Disaccharides and Other Glycosides 237

ch 3

/V-Acetyl-u-D-galactosamine

tCOOH

/V-Acetyl-u-D-neuraminic acid

H— 8 C— OH
9 CH 2 OH

A/-Acetyl-D-neuraminic acid
(open-chain form)

▲ Figure 8.16

can react with methanol in an acidic solution to form an acetal (Figure 8.19). Com- structures of several amino sugars. The amino

pounds containing glycosidic bonds are called glycosides; if glucose supplies the and acetylamino groups are shown in red.

anomeric carbon, they are specifically termed glue os ides. The glycosides include disac-
charides, polysaccharides, and some carbohydrate derivatives.

A. Structures of Disaccharides

Disaccharides are formed when the anomeric carbon of one sugar molecule interacts
with one of several hydroxyl groups in the other sugar molecule. For disaccharides and
other carbohydrate polymers, we must note both the types of monosaccharide residues
that are present and the atoms that form the glycosidic bonds. In the systematic descrip-
tion of a disaccharide we must specify the linking atoms, the configuration of the glyco-
sidic bond, and the name of each monosaccharide residue (including its designation as
a pyranose or furanose). Figure 8.20 presents the structures and nomenclature for four
common disaccharides.

Maltose (Figure 8.20a) is a disaccharide released during the hydrolysis of starch,
which is a polymer of glucose residues. It is present in malt, a mixture obtained from
corn or grain that is used in malted milk and in brewing. Maltose is composed of two D-
glucose residues joined by an a-glycosidic bond. The glycosidic bond links C-l of one
residue (on the left in Figure 8.20a) to the oxygen atom attached to C-4 of the second
residue (on the right). Maltose is therefore a-D-glucopyranosyl-(l — > 4)-D-glucose.

Note that the glucose residue on the left, whose anomeric carbon is involved in the gly-
cosidic bond, is fixed in the a configuration, whereas the glucose residue on the right
(the reducing end, as explained in Section 8.5B) freely equilibrates among the a, /3, and
open-chain structures. (The open-chain form is present in very small amounts). The
structure shown in Figure 8.20a is the /3-pyranose anomer of maltose (the anomer
whose reducing end is in the /3 configuration, the predominant anomeric form).

Cellobiose [/3-D-glucopyranosyl-(l — » 4)-D-glucose] is another glucose dimer
(Figure 8.20b). Cellobiose is the repeating disaccharide in the structure of cellulose, a

CH 2 OH
HO — C — H

ch 2 oh

Glycerol

OH OH

CH 2 OH
H — C — OH

H — C— OH

ch 2 oh

D-Ribitol

◄ Figure 8.17

Structures of several sugar alcohols. Glycerol
(a reduced form of glyceraldehyde) and myo-
inositol (metabolically derived from glucose)
are important constituents of many lipids.
Ribitol (a reduced form of ribose) is a
constituent of the vitamin riboflavin and
its coenzymes.

238 CHAPTER 8 Carbohydrates

(a)

©r

H— 2 C— OH

HO— 3 C — H

H— 4 C— OH

H— 5 C— OH

6 ch 2 oh

D-Gluconate
(open-chain form)

D-Glucono-5-lactone

D-Glucuronate D-Glucuronate

(open-chain form) (/3 pyranose anomer)

▲ Figure 8.18

Structures of sugar acids derived from
o-glucose. (a) Gluconate and its 5-lactone,
(b) The open-chain and pyranose forms
of glucuronate.

plant polysaccharide, and is released during cellulose degradation. The only difference
between cellobiose and maltose is that the glycosidic linkage in cellobiose is (3 (it is a in
maltose). The glucose residue on the right in Figure 8.20b, like the residue on the right
in Figure 8.20a, equilibrates among the a, /3, and open-chain structures.

Lactose [/3-D-galactopyranosyl-(l — » 4)-D-glucose], a major carbohydrate in milk,
is a disaccharide synthesized only in lactating mammary glands (Figure 8.20c). Note
that lactose is an epimer of cellobiose. The naturally occurring a anomer of lactose is
sweeter and more soluble than the (3 anomer. The (3 anomer can be found in stale ice cream,
where it has crystallized during storage and given a gritty texture to the ice cream.

Sucrose [a-D-glucopyranosyl-(l — » 2)-/3-D-fructofuranoside], or table sugar, is the
most abundant disaccharide found in nature (Figure 8.20d). Sucrose is synthesized only in
plants. Sucrose is distinguished from the other three disaccharides in Figure 8.20 because
its glycosidic bond links the anomeric carbon atoms of two monosaccharide residues.
Therefore, the configurations of both the glucopyranose and fructofuranose residues in
sucrose are fixed, and neither residue is free to equilibrate between a and (3 anomers.

B. Reducing and Nonreducing Sugars

Monosaccharides, and most disaccharides, are hemiacetals with a reactive carbonyl
group. They are readily oxidized to diverse products, a property often used in their analy-
sis. Such carbohydrates, including glucose, maltose, cellobiose, and lactose, are some-
times called reducing sugars. Historically, reducing sugars were detected by their ability

Figure 8.19 ►

Reaction of glucopyranose with methanol
produces a glycoside. In this acid-catalyzed
condensation reaction, the anomeric — OH
group of the hemiacetal is replaced by an
— 0CH 3 group, forming methyl glucoside,
an acetal. The product is a mixture of the
a and ft anomers of methyl glucopyranoside.

u-D-Glucopyranose Methanol

Methyl u-D-glucopyranoside

Methyl /3-D-glucopyranoside

8.5 Disaccharides and Other Glycosides

239

/ 3 anomer of maltose

(a-D-Glucopyranosyl-(1^4)-/3-D-glucopyranose)

/ 3 anomer of cellobiose

(/3-D-Glucopyranosyl-(1^>4)-/3-D-glucopyranose)

(c)

a anomer of lactose

(/3-D-Galactopyranosyl-(1-^4)-a-D-glucopyranose)

Sucrose

(a-D-Glucopyranosyl-(1^2)-/3-D-fructofuranoside)

to reduce metal ions such as Cu® or Ag® to insoluble products. Carbohydrates that are
not hemiacetals, such as sucrose, are not readily oxidized because both anomeric carbon
atoms are fixed in a glycosidic linkage. These are classified as nonreducing sugars.

The reducing ability of a sugar polymer is of more than analytical interest. The poly-
meric chains of oligosaccharides and polysaccharides show directionality based on their
reducing and nonreducing ends. There is usually one reducing end (the residue contain-
ing the free anomeric carbon) and one nonreducing end in a linear polymer. All the in-
ternal glycosidic bonds of a polysaccharide involve acetals. The internal residues are not
in equilibrium with open-chain forms and thus cannot reduce metal ions. A branched
polysaccharide has a number of nonreducing ends but only one reducing end.

C. Nucleosides and Other Glycosides

The anomeric carbons of sugars form glycosidic linkages not only with other sugars but
also with a variety of alcohols, amines, and thiols. The most commonly encountered gly-
cosides, other than oligosaccharides and polysaccharides, are the nucleosides, in which a
purine or pyrimidine is attached by its secondary amino group to a /3-D-ribofuranose or
/3-D-deoxyribofuranose moiety. Nucleosides are called N-glycosides because a nitrogen
atom participates in the glycosidic linkage. Guanosine (/3-D-ribofuranosylguanine) is a
typical nucleoside (Figure 8.21). We have already discussed ATP and other nucleotides
that are metabolite coenzymes (Section 7.3). NAD and FAD also are nucleotides.

Two other examples of naturally occurring glycosides are shown in Figure 8.21.
Vanillin glucoside (Figure 8.21b) is the flavored compound in natural vanilla extract.
/3-Galactosides constitute an abundant class of glycosides. In these compounds, a variety
of nonsugar molecules are joined in (3 linkage to galactose. For example, galactocerebro-
sides (see Section 9.5) are glycolipids common in eukaryotic cell membranes and can be
hydrolyzed readily by the action of enzymes called /3-galactosidases.

▲ Figure 8.20

Structures of (a) maltose, (b) cellobiose,

▲ Sugar cane is a major source of commercial
sucrose.

There is a more complete discussion
of nucleosides and nucleotides in
Chapter 19.

240 CHAPTER 8 Carbohydrates

BOX 8.1 THE PROBLEM WITH CATS

(Splenda®), and aspartame (NutraSweet®) bind to the sweet-
ness receptor and cause the sensation of sweetness. They are
hundreds of times more sweet than sucrose.

The sweetness receptor is encoded by two genes called
Taslr2 and Taslr3. We don’t know how sucrose and the other
ligands bind to this receptor even though this is a very active
area of research. In the case of sucrose and the artifical sweet-
eners, how can such different molecules elicit the taste of
sweet?

▲ Cats are carnivores. They probably can’t
taste sweetness.

8.6 Polysaccharides

Polysaccharides are frequently divided into two broad classes. Homoglycans, or ho-
mopolysaccharides, are polymers containing residues of only one type of monosaccha-
ride. Heteroglycans, or heteropolysaccharides, are polymers containing residues of
more than one type of monosaccharide. Polysaccharides are created without a tem-
plate by the addition of particular monosaccharide and oligosaccharide residues. As a
result, the lengths and compositions of polysaccharide molecules may vary within
a population of these molecules. Some common polysaccharides and their structures
are listed in Table 8.2.

Most polysaccharides can also be classified according to their biological roles. For
example, starch and glycogen are storage polysaccharides while cellulose and chitin are
structural polysaccharides. We will see additional examples of the variety and versatility
of carbohydrates when we discuss the heteroglycans in the next section.”

A. Starch and Glycogen

D-Glucose is synthesized in all species. Excess glucose can be broken down to produce
metabolic energy. Glucose residues are stored as polysaccharides until they are needed for
energy production. The most common storage homoglycan of glucose in plants and fungi
is starch and in animals it is glycogen. Both types of polysaccharides occur in bacteria.

8.6 Polysaccharides 241

Table 8.2 Structures of some common polysaccharides

Polysaccharide"

Components)*

Linkage(s)

Storage homoglycans

Starch

Amylose

Glc

a-( 1 —* 4)

Amylopectin

Glc

a-(1 — » 4), a-(1 — » 6) (branches)

Glycogen

Structural homoglycans

Glc

a-(1 — » 4), cx- ( 1 — >6) (branches)

Cellulose

Glc

0 ( 1 - 4 )

Chitin

GIcNAc

0(1-4)

Heteroglycans

Glycosaminoglycans

Disaccharides

(amino sugars, sugar acids)

Various

Hyaluronic acid

GlcUA and GIcNAc

0(1 -3), 0(1 -*4)

°Polysaccharides are unbranched unless otherwise indicated.
fa Glc, Glucose; GIcNAc, N-acetylglucosamine; GlclIA, D-glucuronate.

Guanosine

Vanillin /3-D-glucoside

Starch is present in plant cells as a mixture of amylose and amylopectin and is
stored in granules whose diameters range from 3 to 100 /mm. Amylose is an unbranched
polymer of about 100 to 1000 D-glucose residues connected by a-(l — » 4) glycosidic
linkages, specifically termed a- (l — > 4) glucosidic bonds because the anomeric carbons
belong to glucose residues (Figure 8.22a). The same type of linkage connects glucose
monomers in the disaccharide maltose (Figure 8.20a). Although it is not truly soluble in
water, amylose forms hydrated micelles in water and can assume a helical structure
under some conditions (Figure 8.22b).

Amylopectin is a branched version of amylose (Figure 8.23). Branches, or poly-
meric side chains, are attached via a-(l — >6) glucosidic bonds to linear chains of
residues linked by a- (l — » 4) glucosidic bonds. Branching occurs, on average, once
every 25 residues and the side chains contain about 15 to 25 glucose residues. Some side
chains themselves are branched. Amylopectin molecules isolated from living cells may
contain 300 to 6000 glucose residues.

An adult human consumes about 300 g of carbohydrate daily, much of which is in
the form of starch. Raw starch granules resist enzymatic hydrolysis but cooking causes
them to absorb water and swell. The swollen starch is a substrate for two different gly-
cosidases. Dietary starch is degraded in the gastrointestinal tract by the actions of a-
amylase and a debranching enzyme, a- Amylase, which is present in both animals and

/3-D-Galactosyl 1 -glycerol
▲ Figure 8.21

Structures of three glycosides. The nonsugar
components are shown in blue, (a) Guano-
sine. (b) Vanillin glucoside, the flavored com-
pound in vanilla extract, (c) /3-D-Galactosyl
1-glycerol, derivatives of which are common
in eukaryotic cell membranes.

Starch metabolism is described in
Chapter 15.

(a)

(b)

CH 2 OH

CH 7 OH

CH 2 OH

▲ Figure 8.22

Amylose. (a) Structure of amylose. Amylose, one form of starch, is a linear polymer of glucose
residues linked by a-( 1 -^4)-D-glucosidic bonds, (b) Amylose can assume a left-handed helical
conformation, which is hydrated on the inside as well as on the outer surface.

242 CHAPTER 8 Carbohydrates

Figure 8.23 ►

Structure of amylopectin. Amylopectin, a
second form of starch, is a branched polymer.
The linear glucose residues of the main
chain and the side chains of amylopectin
are linked by a- (l -^4)-D-glucosidic bonds,
and the side chains are linked to the main
chain by a-( 1 —> 6)-D-glucosidic bonds.

'W\/'

plants, is an endoglycosidase (it acts on internal glycosidic bonds). The enzyme catalyzes
random hydrolysis of the a- (l — » 4) glucosidic bonds of amylose and amylopectin.

Another hydrolase, /3- amylase, is found in the seeds and tubers of some plants.
/3- Amylase is an exoglycosidase (it acts on terminal glycosidic bonds). It catalyzes se-
quential hydrolytic release of maltose from the free, nonreducing ends of amylopectin.

Despite their a and /3 designations, both types of amylases act only ona-(1^4)-D-
glycosidic bonds. Figure 8.24 shows the action of a- amylase and /3-amylase on amy-
lopectin. The a- (l — > 6) linkages at branch points are not substrates for either a- or
/3- amylase. After amylase-catalyzed hydrolysis of amylopectin, highly branched cores re-
sistant to further hydrolysis, called limit dextrins, remain. Limit dextrins can be further
degraded only after debranching enzymes have catalyzed hydrolysis of the a- (l —> 6)
linkages at branch points.

Glycogen is also a branched polymer of glucose residues. Glycogen contains the
same types of linkages found in amylopectin but the branches in glycogen are smaller
and more frequent, occurring every 8-12 residues. In general, glycogen molecules are
larger than starch molecules, Glycogen up to contains 50,000 glucose residues. In mammals,

Figure 8.24 ►

Action of a-amylase and /3-amylase on
amylopectin. a-Amylase catalyzes random
hydrolysis of internal a-( 1 — >4) glucosidic
bonds; /3-amylase acts on the nonreducing
ends. Each hexagon represents a glucose
residue; the single reducing end of the
branched polymer is red. (An actual amy-
lopectin molecule contains many more
glucose residues than shown here.)

8.6 Polysaccharides 243

depending on the nutritional state, glycogen can account for up to 10% of the mass of
the liver and 2% of the mass of muscle.

The branched structures of amylopectin and glycogen possess only one reducing
end but many nonreducing ends. The reducing end of glycogen is covalently attached to
a protein called glycogenin (Section 12. 5 A). Enzymatic lengthening and degradation of
polysaccharide chains occurs at the nonreducing ends.

Enzymes that catalyze the intracellular
synthesis and breakdown of glycogen
are described in Chapter 12.

B. Cellulose

Cellulose is a structural polysaccharide. It is a major component of the rigid cell walls
that surround many plant cells. The stems and branches of many plants consist largely
of cellulose. This single polysaccharide accounts for a significant percentage of all or-
ganic matter on Earth. Like amylose, cellulose is a linear polymer of glucose residues,
but in cellulose the glucose residues are joined by /?-( 1 — >4) linkages rather than
a-( 1 — > 4) linkages. The two glucose residues of the disaccharide cellobiose also are
connected by a /3-( 1 — » 4) linkage (Figure 8.20b). Cellulose molecules vary greatly in
size, ranging from about 300 to more than 15,000 glucose residues.

The (3 linkages of cellulose result in a rigid extended conformation in which each
glucose residue is rotated 180° relative to its neighbors (Figure 8.25). Extensive hydro-
gen bonding within and between cellulose chains leads to the formation of bundles, or
fibrils (Figure 8.26). Cellulose fibrils are insoluble in water and are quite strong and
rigid. Cotton fibers are almost entirely cellulose and wood is about half cellulose. Be-
cause of its strength, cellulose is used for a variety of purposes and is a component of a
number of synthetic materials including cellophane and the fabric rayon. We are most
familiar with cellulose as the main component of paper.

Enzymes that catalyze the hydrolysis of a-D-glucosidic bonds (a-glucosidases, such
as a- and /3- amylase) do not catalyze the hydrolysis of /3-D-glucosidic bonds. Similarly,
/3-glucosidases (such as cellulase) do not catalyze the hydrolysis of a-D-glucosidic
bonds. Humans and other mammals can metabolize starch, glycogen, lactose, and su-
crose and use the monosaccharide products in a variety of metabolic pathways. Mam-
mals cannot metabolize cellulose because they lack enzymes capable of catalyzing the
hydrolysis of /3-glucosidic linkages. Ruminants such as cows and sheep have microor-
ganisms in their rumen (a compartment in their multichambered stomachs) that
produce /3-glucosidases. Thus, ruminants can obtain glucose from grass and other
plants that are rich in cellulose. Because they have cellulase-producing bacteria in their
digestive tracts, termites also can obtain glucose from dietary cellulose.

(a)

▲ Figure 8.25

Structure of cellulose. Note the alternating orientation of successive glucose residues in the cellu-
lose chain, (a) Chair conformation, (b) Modified Haworth projection.

▲ Figure 8.26

Cellulose fibrils. Intra-and interchain hydro-
gen bonding gives cellulose its strength and
rigidity.

244 CHAPTER 8 Carbohydrates

Figure 8.27 ►

Structure of chitin. The linear homoglycan
chitin consists of repeating units of
P~( 1 — > 4)-l i nked GIcNAc residues. Each
residue is rotated 180° relative to its
neighbors.

▲ The giant redwood trees of California con-
tains tons of cellulose.

▲ Cellulose fibers. Plants make large cellu-
lose fibers that serve as structural support.
A scanning electron micrograph of these
fibers shows how they overlap to form a
large net-like sheet. These cellulose fibers
are about 253 million years old. They were
recovered from deep within a salt mine in
New Mexico.

C=0

ch 3 ch 3

C. Chitin

Chitin, probably the second most abundant organic compound on Earth, is a structural
homoglycan found in the exoskeletons of insects and crustaceans and also in the cell
walls of most fungi and red algae. Chitin is a linear polymer similar to cellulose. It is
made up of /3-( 1 —> 4) -linked GIcNAc residues rather than glucose residues (Figure 8.27).
Each GIcNAc residue is rotated 180° relative to its neighbors. The GIcNAc residues in
adjacent strands of chitin form hydrogen bonds with each other resulting in linear
fibrils of great strength. Chitin is often closely associated with nonpolysaccharide
compounds, such as proteins and inorganic material.

8.7 Glycoconjugates

Glycoconjugates consist of polysaccharides linked to (conjugated with) proteins or
peptides. In most cases, the polysaccharides are composed of several different mono-
saccharide units. Thus, they are heteroglycans. (Starch, glycogen, cellulose, and chitin are
homoglycans.) Heteroglycans appear in three types of glycoconjugates — proteoglycans,
peptidoglycans, and glycoproteins. In this section, we see how the chemical and physi-
cal properties of the heteroglycans in glycoconjugates are suited to various biological
functions.

A. Proteoglycans

Proteoglycans are complexes of proteins and a class of polysaccharides called glycos-
aminoglycans. These glycoconjugates occur predominately in the extracellular matrix
(connective tissue) of multicellular animals.

Glycosaminoglycans are unbranched heteroglycans of repeating disaccharide units.
As the name gly cos amino gly can indicates, one component of the disaccharide is an
amino sugar, either D-galactosamine (GalN) or D-glucosamine (GlcN). The amino
group of the amino -sugar component can be acetylated forming N- acetylgalactosamine
(GalNAc) or GIcNAc. The other component of the repeating disaccharide is usually an
alduronic acid. Specific hydroxyl and amino groups of many glycosaminoglycans are
sulfated. These sulfate groups and the carboxylate groups of alduronic acids make gly-
cosaminoglycans polyanionic.

Several types of glycosaminoglycans have been isolated and characterized. Each
type has its own sugar composition, linkages, tissue distribution, and function and each
is attached to a characteristic protein. Hyaluronic acid is an example of a glycosamino-
glycan composed of the repeating disaccharide unit shown in Figure 8.28. It is found in
the fluid of joints where it forms a viscous solution that is an excellent lubricant.
Hyaluronic acid is also a major component of cartilage.

Up to 100 glycosaminoglycan chains can be attached to the protein of a proteogly-
can. These heteroglycan chains are usually covalently bound by a glycosidic linkage to

8.7 Glycoconjugates 245

◄ Figure 8.28

Structure of the repeating disaccharide of
hyaluronic acid. The repeating disaccharide
of this glycosaminoglycan contains D-glu-
curonate (GlcLIA) and GIcNAc. Each GlcLIA
residue is linked to a GIcNAc residue
through p-( 1 — >3) linkage; each GIcNAc
residue is in turn linked to the next GlcLIA
residue through a /3-(l — >4) linkage.

the hydroxyl oxygens of serine residues. (Not all glycosaminoglycans are covalently
linked to proteins.) Glycosaminoglycans can account for up to 95% of the mass of a
proteoglycan.

Proteoglycans are highly hydrated and occupy a large volume because their gly-
cosaminoglycan component contains polar and ionic groups. These features confer
elasticity and resistance to compression — important properties of connective tissue. For
example, the flexibility of cartilage allows it to absorb shocks. Some of the water can be
pressed out when cartilage is compressed but relief from pressure allows cartilage to re-
hydrate. In addition to maintaining the shapes of tissues, proteoglycans can also act as
extracellular sieves and help direct cell growth and migration.

Examination of the structure of cartilage shows how proteoglycans are organized
in this tissue. Cartilage is a mesh of collagen fibers (Section 4.1 1) interspersed with large
proteoglycan aggregates (M r ~2 x 10 8 ). Each aggregate assumes a characteristic shape
that resembles a bottle brush (Figure 8.29). These aggregates contain hyaluronic acid
and several other glycosaminoglycans, as well as two types of proteins — core proteins
and link proteins. A central strand of hyaluronic acid runs through the aggregate and
many proteoglycans — core proteins with glycosaminoglycan chains attached — branch
from its sides. The core proteins interact noncovalently with the hyaluronic acid
strand, mostly by electrostatic interactions. Link proteins stabilize the core protein-
hyaluronic acid interactions.

The major proteoglycan of cartilage is called aggrecan. The protein core of aggrecan
(M r ~ 220,000) carries approximately 30 molecules of keratan sulfate (a glycosamino-
glycan composed chiefly of alternating N-acetylglucosamine 6-sulfate and galactose
residues) and approximately 100 molecules of chondroitin sulfate (a glycosaminoglycan

Proteoglycans (core
proteins with

glycosaminoglycan Central strand of

chains attached) hyaluronic acid

▲ Lobsters have an exoskeleton made of chitin.

The color of the exoskeleton is determined
by the foods that the lobster eats. When
it ingests p- carotene derivatives they are
converted to a complex mixture of protein-
bound carotenes called crustacayanin
that has a greenish-brown color. When
lobsters are cooked, the crustacyanin breaks
down, releasing free /1-carotene derivatives
that are red in color, like the red color of
maple leaves in autumn (see Section 15.1).

◄ Figure 8.29

Proteoglycan aggregate of cartilage. Core pro-
teins carrying glycosaminoglycan chains are
associated with a central strand of a single
hyaluronic acid molecule. These proteins
have many covalently attached glycosamino-
glycan chains (keratan sulfate and chon-
droitin sulfate molecules). The interactions
of the core proteins with hyaluronic acid are
stabilized by link proteins, which interact
noncovalently with both types of molecules.
The aggregate has the appearance of a bottle
brush.

246 CHAPTER 8 Carbohydrates

BOX 8.2 NODULATION FACTORS ARE LIPO-OLIGOSACCHARIDES

Legumes such as alfalfa, peas, and soybeans develop organs
called nodules on their roots. Certain soil bacteria (rhizobia)
infect the nodules and, in a symbiosis with the plants, carry
out nitrogen fixation (reduction of atmospheric nitrogen to
ammonia). The symbiosis is highly species-specific: only cer-
tain combinations of legumes and bacteria can cooperate and
therefore these organisms must recognize each other. Rhizobia
produce extracellular signal molecules that are oligosaccha-
rides called nodulation factors. Extremely low concentrations
of these compounds can induce their plant hosts to develop
the nodules that the rhizobia can infect. A host plant responds
only to a nodulation factor of a characteristic composition.

Infection begins when the plant root hair recognizes the
nodulation factor via surface Nod-factor receptors. This results
in a response that allows the bacteria to enter the root hair and
migrate down to the cells in the root where the nodule forms.

All the nodulation factors studied to date are oligosac-
charides that have a linear chain of /3-(l —> 4) N-acetylglu-
cosamine (GlcNAc) — the same repeating structure as in
chitin (Section 8.6b). Most nodulation factors are sugar pen-
tamers although the number of residues can vary between
three and six (see figure below). Species specificity is pro-
vided by variation in polymer length and potential substitu-
tion on five sites at the nonreducing end (R1 to R5) and two
sites at the reducing end (R6 and R7). Rl, an acyl group sub-
stituting the nitrogen atom at C-2 of the nonreducing end, is
a fatty acid, usually 18 carbons long. Thus, the nodulation
factors are lipo-oligosaccharides. R6, bound to the alcohol at
C-6 of the reducing end, can have a wide variety of struc-
tures, including sulfate or methyl fucose. Research on these
growth regulators for legumes has stimulated the search for
biological activities of other oligosaccharides.

See Section 17.1 for details about
nitrogen fixation.

▲ General structure of nodulation factors, lipo-oligosaccharides with an
/V-acetylglucosamine (GlcNAc) backbone. The number of internal residues of
A/-acetylglucosamine is shown by n, which is usually 3 but can sometimes
be 1, 2, or 4. Rl is a fatty acyl substituent, usually 18 carbons long.

▲ Formation of nodules in the legume Lotus
japonicus. Rhizobia (blue) have secreted nodula-
tion factor leading to endocytosis by root hair
cells and formation of an infection thread con-
necting the point of uptake (top) to the root
nodule cells (below).

composed of alternating N-acetylgalactosamine sulfate and glucuronate residues). Ag-
grecan is a member of a small family of hyalectans, proteoglycans that bind to
hyaluronic acid. Other hyalectans provide elasticity to blood vessel walls and modulate
cell-cell interactions in the brain.

B. Peptidoglycans

Peptidoglycans are polysaccharides linked to small peptides. The cell walls of many bac-
teria contain a special class of peptidoglycan with a heteroglycan component attached
to a four or five residue peptide. The heteroglycan component is composed of alternating
residues of GlcNAc and N-acetylmuramic acid (MurNAc) joined by /3-(l — > 4) link-
ages (Figure 8.30). MurNAc is a nine-carbon sugar found only in bacteria. MurNAc
consists of the three-carbon acid D-lactate joined by an ether linkage to C-3 of GlcNAc.

8.7 Glycoconjugates 247

oh 3

◄ Figure 8.30

Structure of the polysaccharide in bacterial cell

C = 0
1

wall peptidoglycan. The glycan is a polymer
of alternating GIcNAc and A/-acetyl mu ramie

ch 2 oh

acid (MurNAc) residues.

GIcNAc

The polysaccharide moiety of peptidoglycans resembles chitin except that every second
GIcNAc residue is modified by addition of lactate to form MurNAc. The antibacterial
action of lysozyme (Section 6.6) results from its ability to catalyze hydrolysis of the
polysaccharide chains of peptidoglycans.

The peptide component of peptidoglycans varies among bacteria. The peptide
component in Staphylococcus aureus is a tetrapeptide with alternating l and d amino
acids: L-Ala-D-Isoglu-L-Lys-D-Ala. (Isoglu represents isoglutamate, a form of gluta-
mate in which the y-c arboxyl group — not the a-carboxyl group — is linked to the next
residue.) Other species have a different amino acid at the third position. An amide bond
links the amino group of the L-alanine residue to the lactyl carboxylate group of a
MurNAc residue of the glycan polymer (Figure 8.31). The tetrapeptide is cross-linked to
another tetrapeptide on a neighboring peptidoglycan molecule by a chain of five glycine
residues (pentaglycine). Pentaglycine joins the L-lysine residue of one tetrapeptide to
the carboxyl group of the D-alanine residue of the other tetrapeptide. Extensive cross-
linking essentially converts the peptidoglycan to one huge, rigid, macromolecule that
defines the shape of the bacterium by covering its plasma membrane and protecting the
cell from fluctuations in osmotic pressure.

Most bacteria have an additional exterior layer of dense polysaccharide called the
capsule. The capsule is made up of chains of polysaccharide composed mainly of
N-acetylglucosamine (GIcNAc) residues but various other amino sugars are present.
The capsule protects the bacterial cell from injury. The capsule in pathogenic bacteria
help cells avoid destruction by the immune system.

In gram-negative bacteria, the peptidoglycan cell wall lies between the inner plasma
membrane and the outer membrane. In gram-positive bacteria, there is no outer mem-
brane and the cell wall is much thicker. This is one of the reasons why the Gram stain
(named after Christian Gram) will color the surfaces of some bacteria (gram positive)
and not others (gram negative).

During peptidoglycan biosynthesis, a five-residue peptide — L-Ala-D-Isoglu-L-
Lys-D-Ala-D-Ala — is attached to a MurNAc residue. In subsequent steps, five glycine
residues are added sequentially to the £- amino group of the lysine residue forming the
pentaglycine bridge. In the final step of synthesis, a transpeptidase catalyzes formation
of a peptide linkage between the penultimate alanine residue and a terminal glycine
residue of a pentaglycine bridge of a neighboring peptidoglycan strand. This reaction is
driven by release of the terminal D-alanine residue.

The structure of the antibiotic penicillin (Figure 8.32) resembles the terminal
D-Ala-D-Ala residues of the immature peptidoglycan. Penicillin binds, probably irre-
versibly, to the transpeptidase active site inhibiting the activity of the enzyme and
thereby blocking further peptidoglycan synthesis. The antibiotic prevents growth and
proliferation of bacteria. Penicillin is selectively toxic to bacteria because the reaction it
affects occurs only in certain bacteria, not in eukaryotic cells.

▲ Staphylococcus aureus cells. These bacter-
ial cells have extensive polysaccharide
capsules that protect them from their host’s
immune system.

▲ The Gram stain. The Gram staining proce-
dure distinguishes between gram-positive
bacteria (left, purple) and gram-negative
bacteria (right, pink).

248 CHAPTER 8 Carbohydrates

(a) MurNAc GIcNAc CH 3

C=0

(b) Polysaccharide

L-Alanine

D-lsoglutamate

CH — CH 3

c=o

: i

1 0

CH — COO u

(CH 2 ) 2

▲ Figure 8.31

Structure of the peptidoglycan of Staphylococcus aureus, (a) Repeating disaccharide unit, tetrapeptide,
and pentaglycine components. The tetrapeptide (blue) is linked to a MurNAc residue of the glycan
moiety (black). The e-amino group of the L-lysine residue of one tetrapeptide is cross-linked to the
a-carboxyl group of the D-alanine residue of another tetrapeptide on a neighboring peptidoglycan
molecule via a pentaglycine bridge (red), (b) Cross-linking of the peptidoglycan macromolecule.

C=0

NH OOOOO

L-Lysine CH — (CH 2 ) 4 — N — C — CH 2 — N — C — CH 2 — N — C — CH 2 — N — C — CH 2 — N — C — CH 2 — N —

H H H H H

c=o 1 1

Pentaglycine bridge

D-Alanine

CH — CH 3

COO

-c-

-N-

H^S X /

-CHq

CH 3

COO

II H /CH 3

C — N — C

H I H

</” N “

ch 3

-CH

COO

C. Glycoproteins

Glycoproteins, like proteoglycans, are proteins that contain covalently bound oligosac-
charides (i.e., proteins that are glycosylated). In fact, proteoglycans are a type of glyco-
protein. The carbohydrate chains of a glycoprotein vary in length from one to more
than 30 residues and can account for as much as 80% of the total mass of the molecule.
Glycoproteins are an extraordinarily diverse group of proteins that includes enzymes,
hormones, structural proteins, and transport proteins.

The oligosaccharide chains of different glycoproteins exhibit great variability in com-
position. The composition of oligosaccharide chains can vary even among molecules of
the same protein, a phenomenon called microheterogeneity.

Several factors contribute to the structural diversity of the oligosaccharide chains of
glycoproteins.

1. An oligosaccharide chain can contain several different sugars. Eight sugars predomi-
nate in eukaryotic glycoproteins: the hexoses L-fucose, D-galactose, D-glucose, and
D-mannose; the hexosamines N-acetyl-D-galactosamine and N-acetyl-D-glucosamine;
the nine-carbon sialic acids (usually N-acetylneuraminic acid); and the pentose
D-xylose. Many different combinations of these sugars are possible.

2. The sugars can be joined by either a- or /J-glycosidic linkages.

v/wv/'D-Ala-D-Ala

▲ Figure 8.32

Structures of penicillin and -D-Ala-D-Ala. The

portion of penicillin that resembles the
dipeptide is shown in red. R can be a variety
of substituents.

3. The linkages can also join various carbon atoms in the sugars. In hexoses and hex-
osamines, the glycosidic linkages always involve C- 1 of one sugar but can involve C-2,
C-3, C-4, or C-6 of another hexose or C-3, C-4, or C-6 of an amino sugar (C-2 is
usually N-acetylated in this class of sugar). C-2 of sialic acid, not C-l, is linked to
other sugars.

4. Oligosaccharide chains of glycoproteins can contain up to four branches.

8.7 Glycoconjugates 249

(a)

ch 3

(b)

o c

c=o

ch 3

Asparagine

residue

◄ Figure 8.33

0-Glycosidic and /V-glycosidic linkages.

(a) /V-Acetylgalactosamine-serine linkage, the
major O-glycosidic linkage found in glycopro-
teins. (b) /V-Acetylglucosamine-asparagine
linkage, which characterizes N-linked glyco-
proteins. The O-glycosidic linkage is a,
whereas the A/-glycosidic linkage is p.

The astronomical number of possible oligosaccharide structures afforded by these
four factors is not realized in cells because cells do not possess specific glycosyltrans-
ferases to catalyze the formation of all possible glycosidic linkages. In addition, individual
glycoproteins — through their unique conformations — modulate their own interactions
with the glycosylating enzymes so that most glycoproteins possess a heterogeneous but
reproducible oligosaccharide structure.

The oligosaccharide chains of most glycoproteins are either O- or N-linked. In
0-linked oligosaccharides, a GalNAc residue is typically linked to the side chain of a ser-
ine or threonine residue. In W-linked oligosaccharides, a GlcNAc residue is linked to the
amide nitrogen of an asparagine residue. The structures of an O-glycosidic and an
N-glycosidic linkage are compared in Figure 8.33. Additional sugar residues can be
attached to the GalNAc or the GlcNAc residue. An individual glycoprotein can contain
both O- and N-linked oligosaccharides and some glycoproteins contain a third type of
linkage. In these glycoproteins, the protein is attached to ethanolamine that is linked to
a branched oligosaccharide to which lipid is also attached (Section 9.10).

There are four important subclasses of O-glycosidic linkages in glycoproteins.

1. The most common O-glycosidic linkage is the GalNAc- Ser/Thr linkage mentioned
above. Other sugars — for example, galactose and sialic acid — are frequently linked
to the GalNAc residue (Figure 8.34a).

2. Some of the 5-hydroxylysine (Hyl) residues of collagen (Figure 4.35) are joined to
D-galactose via an O-glycosidic linkage (Figure 8.34b). This structure is unique to
collagen.

3. The glycosaminoglycans of certain proteoglycans are joined to the core protein via
a Gal-Gal-Xyl-Ser structure (Figure 8.34c).

4. In some proteins, a single residue of GlcNAc is linked to serine or threonine
(Figure 8.34d).

(a)

NeuNAc a-(2 — > 3) GalNAc /3-(l -^3)

^GalNAc — er/Thr
NeuNAc a-( 2^6)/

(b)

(c)

(d)

— Gal— lyl

— Gal — Gal — Xyl — >er

GlcNAc — Ser/Thr

◄ Figure 8.34

Four subclasses of O-glycosidic linkages.

(a) Example of a typical linkage in which N-
acetylgalactosamine (GalNAc) with attached
residues is linked to a serine or threonine
residue, (b) Linkage found in collagen,
where a galactose residue, usually attached
to a glucose residue, is linked to hydroxyly-
sine (Hyl). (c) Trisaccharide linkage found in
certain proteoglycans, (d) GlcNAc linkage
found in some proteins.

250 CHAPTER 8 Carbohydrates

BOX 8.3 ABO BLOOD GROUP

The ABO blood group was first discovered in 1901 by Karl
Landsteiner, who received the Nobel Prize in Physiology or
Medicine in 1930. Most primates display three different kinds
of O- or N-linked oligosaccharides on their cell surfaces. The
core structure of these oligosaccharides is called H antigen. It
consists of various combinations of galactose (Gal), fucose
(Fuc), N-acetylglucosamine (GlcNac), and N-acetylneuraminic
acid (sialic acid, NeuNAc). These monosaccharides are linked
in various ways to form a short branched structure that ex-
hibits considerable microheterogeneity. One of the most com-
mon H antigen structures is shown in the figure.

The core structure (H antigen) can be modified in various
ways. The addition of a GalNAc residue in a- (l — > 3) linkage
forms A antigen. This reaction is catalyzed by A enzyme. The
addition of Gal in a- (l — > 3) linkage is catalyzed by B enzyme.

If only A antigen is present, a person will have A blood
type. If only B antigen is present, the blood type will be B.
The AB blood type indicates that both A antigen and B anti-

gen are present on cell surfaces. If neither GalNAc or Gal
have been added to the H antigen structure, then neither A
antigen nor B antigen will be present and the blood type is O.

The ABO blood group is determined by a single gene on
chromosome 9. Human (and other primate) populations
contain many alleles of this gene. The original gene encoded
A enzyme, which transfers GalNAc. Variants of this gene have
altered the specificity of the enzyme so that it no longer rec-
ognizes GalNAc but, instead, transfers Gal. These B enzymes
differ by several amino acid residues from the allele that en-
codes the A enzyme. The structures of both types of glycosyl-
transferase enzymes have been solved and they reveal that
only a single amino acid substitution is required to change
the specificity from iV-acetylaminogalactosyltransferase to
galactosyltransferase.

The chromosome 9 locus can also contain several alleles
that encode nonfunctional proteins. One of the most common
mutations is a single base pair deletion near the N-terminus

FUC a-(1 — >2) Hanti9en

Gal /3-(1 3)- GIcNAc /3...

A enzyme^^ B enzyme

Fuc a-{ 1 ->2) Fuc a-( 1 -^2)

\ \

Gal j3-(1-» 3)- GIcNAc p... Gal /3-(1 3)- GIcNAc /3...

GalNAc a-0 Gal a-{ 1^3^

A antigen B antigen

O-Linked oligosaccharides may account for 80% of the mass of mucins. These
large glycoproteins are found in mucus, the viscous fluid that protects and lubricates
the epithelium of the gastrointestinal, genitourinary, and respiratory tracts. The
oligosaccharide chains of mucins contain an abundance of NeuNAc residues and sul-
fated sugars. The negative charges of these residues are responsible in part for the
extended shape of mucins, which contributes to the viscosity of solutions containing
mucins.

The biosynthesis of the oligosaccharide chains of glycoproteins requires a battery
of specific enzymes in distinct compartments of the cell. In the stepwise synthesis of
O-linked oligosaccharides, glycosyltransferases catalyze the addition of glycosyl groups
donated by nucleotide-sugar coenzymes. The oligosaccharide chains are assembled by
addition of the first sugar molecule to the protein, followed by subsequent single-sugar
additions to the nonreducing end.

N-Linked oligosaccharides, like O-linked oligosaccharides, exhibit great variety in
sugar sequence and composition. Most N-linked oligosaccharides can be divided into

8.7 Glycoconjugates 251

of the coding region. This deletion shifts the reading frame
for translation (Section 22.1) making it impossible to synthe-
size a functional enzyme of either type. This is another ex-
ample of a human pseudogene. People who are homozygous
for these nonfunctional O alleles will not synthesize either A
antigen or B antigen and their blood type will be O. (See the
Online Medelian Inheritance in Man (OMIM: ncbi.nlm.nih.
gov/omim) database entry 1 10300 for an excellent and complete
summary of all ABO variants.)

All of your blood cells display some of the unmodified
core oligosaccharide (H antigen) even if your blood type is A,
B, or AB. This is because not all of the H antigen structures
are modified. Under normal circumstances, human plasma
will not contain antibodies against H antigen. However,
O-type individuals will have antibodies against A antigen and
B antigen because these structures are recognized as nonself.
If O-type individuals receive a blood transfusion from some-
one with A, B, or AB blood, they will mount an immune re-
sponse and reject it. Similarly, if you have A- type blood you
will have anti-B antibodies and cannot receive a transfusion
from someone with B or AB blood type.

The O allele (pseudogene) is the most common allele in
most human populations and the B allele is the most rare.
Some Native American populations are homogeneous for the
O allele and everyone has type O blood. Type O individuals
are perfectly normal, indicating that the absence of the A and
B oligosaccharide structures has no effect on normal growth
and development (i.e., the allele is neutral in most environ-
ments). However, there are some correlations between blood
type and disease. People with type O blood, for example, are
more susceptible to cholera caused by infections of the bac-
terium Vibrio cholerae. Such selective pressures may be re-
sponsible for maintaining the frequencies of A and B alleles
in some populations.

Percent of
population
that has the
O blood type

□ 50-60

□ 60-70

□ 70-80

□ 80-90
H 90-100

▲ ABO blood group: distribution of alleles in humans.

three subclasses: high mannose, complex, and hybrid (Figure 8.35). The appearance of a
common core pentasaccharide (GlcNAc 2 Man 3 ) in each class reflects a common initial
pathway for biosynthesis. The synthesis of N-linked oligosaccharides begins with the as-
sembly of a compound consisting of a branched oligosaccharide with 14 residues (nine
of which are mannose residues) linked to the lipid dolichol. The entire oligosaccharide
chain is transferred to an asparagine residue of a newly synthesized protein, after which
the chain is trimmed by the action of glycosidases. High-mannose chains represent an
early stage in the biosynthesis of N-linked oligosaccharides. Complex oligosaccharide
chains result from further removal of sugar residues from high-mannose chains and the
addition of other sugar residues, such as fucose, galactose, GlcNAc, and sialic acid (a
phenomenon called oligosaccharide processing). These additional sugar residues are
donated by nucleotide sugars in reactions catalyzed by glycosyltransferases as in the
synthesis of O-linked oligosaccharides. In certain cases, a glycoprotein can contain a hy-
brid oligosaccharide chain, a branched oligosaccharide in which one branch is of the
high-mannose type and the other is of the complex type.

252

CHAPTER 8 Carbohydrates

(a)

Man o;-(i — > 2) Man a-( 1^2) Man «-(i -^3)

Man a-(i -> 2) Man a-( 1 -> 3)

Man /3-(l^4) GIcNAc /3-(i-»4) GIcNAc - Asn

Man a-(i -> 2) Man a-( 1 -> 6)'

X M

an «-(i^6)

(b) SA a-(2 ->3,6) Gal jS-(l 4) GIcNAc /3-(l^ 2) Man «-(i -^3)

Man /3-(i-»4) GIcNAc /3-(l->4) GIcNAc

Asn

(c)

▲ Figure 8.35

Structures of /V-linked oligosaccharides, (a)

High-mannose chain, (b) Complex chain.

(c) Hybrid chain. The pentasaccharide core
common to all A/-linked structures is shown
in red. SA represents sialic acid, usually
NeuNAc.

▲ Mucins. Mucins are heavily glycosylated
proteins secreted by the epithelial cells of
animals. You are probably familiar with the
mucins secreted by cells lining your mouth
(saliva), nasal cavity (“snot”), and intestine.
The mucin shown here is being secreted by
a hagfish.

The synthesis of glycoproteins is dis-
cussed in Section 22.10.

Gal /3-(l 4) GIcNAc /3-(l^2) Man a-(l -^3)

Man ck-(1 — > 3)

Man /3-(i — > 4) GIcNAc j3-(i->4) GIcNAc - Asn

Man 0-0^6)'

Man «-(i -> 6)

Most glycoproteins are secreted from the cell or are bound to the outer surface of
the plasma membrane. There are very few glycoproteins in the cytoplasm. With rare ex-
ceptions, none of the basic metabolic enzymes are glycosylated. The addition of
oligosaccharide chains is tightly coupled to sorting and secretion in eukaryotic cells.
The oligosaccharides are attached to specific proteins in the lumen of the endoplasmic
reticulum and the groups are modified by various glycosyltransferase enzymes as the
proteins move from the ER through the Golgi to the cell surface. The structure of the
linked oligosaccharide serves as a marker for sorting proteins into various compart-
ments. For example, some proteins are targeted to the lysosomes, depending on the
structure of the oligosaccharide, while others are marked for secretion.

In addition to their roles as markers in sorting and secretion, the presence of one or
more oligosaccharide chains on a protein can alter its physical properties, including its
size, shape, solubility, electric charge, and stability. Biological properties that can be
altered include rate of secretion, rate of folding, and immunogenicity. In a few cases,
specific roles for the oligosaccharide chains of glycoproteins have been identified. For
example, a number of mammalian hormones are dimeric glycoproteins whose oligosac-
charide chains facilitate assembly of the dimer and confer resistance to proteolysis. Also,
the recognition of one cell by another that occurs during cell migration or oocyte fertil-
ization can depend in part on the binding of proteins on the surface of one cell to the
carbohydrate portions of certain glycoproteins on the surface of the other cell.

Summary

1. Carbohydrates include monosaccharides, oligosaccharides, and
polysaccharides. Monosaccharides are classified as aldoses or ke-
toses or their derivatives.

2. A monosaccharide is designated D or L, according to the configu-
ration of the chiral carbon farthest from the carbonyl carbon
atom. Each monosaccharide has 2” possible stereoisomers,
where n is the number of chiral carbon atoms. Enantiomers are
nonsuperimposable mirror images of each other. Epimers differ
in configuration at only one of several chiral centers.

3. Aldoses with at least five carbon atoms and ketoses with at least
six carbon atoms exist principally as cyclic hemiacetals or
hemiketals known as furanoses and pyranoses. In these ring
structures, the configuration of the anomeric (carbonyl) carbon
is designated either a or (3 . Furanoses and pyranoses can adopt
several conformations.

4. Derivatives of monosaccharides include sugar phosphates, deoxy
sugars, amino sugars, sugar alcohols, and sugar acids.

Problems 253

5. Glycosides are formed when the anomeric carbon of a sugar forms
a glycosidic linkage with another molecule. Glycosides include dis-
accharides, polysaccharides, and some carbohydrate derivatives.

6. Homoglycans are polymers containing a single type of sugar
residue. Examples of homoglycans include the storage polysac-
charides starch and glycogen and the structural polysaccharides
cellulose and chitin.

7. Hetero glycans contain more than one type of sugar residue. They
are found in glycoconjugates such as proteoglycans, peptidogly-
cans, and glycoproteins.

8. Proteoglycans are proteins linked to chains of repeating disaccha-
rides. Proteoglycans are prominent in the extracellular matrix and
in connective tissues such as cartilage.

9. The cell walls of many bacteria are made of peptidoglycans that are
heteroglycan chains linked to peptides. Peptidoglycan molecules
are extensively cross-linked, essentially converting peptidoglycan
into a rigid macromolecule that defines the shape of a bacterium
and protects the plasma membrane.

10. Glycoproteins are proteins containing covalently bound oligosac-
charides. The oligosaccharide chains of most glycoproteins are
either O-linked to serine or threonine residues or TV-linked to
asparagine residues and exhibit great variety in structure and
sugar composition.

Problems

1. Identify each of the following:

(a) Two aldoses whose configuration at carbons 3, 4, and 5
matches that of D-fmctose.

(b) The enantiomer of D-galactose.

(d) A ketose that has no chiral centers.

(e) A ketose that has only one chiral center.

(f) Monosaccharide residues of cellulose, amylose, and glycogen.

(g) Monosaccharide residues of chitin.

2. Draw Fischer projections for (a) L-mannose, (b) L-fucose
(6-deoxy-L-galactose), (c) D-xylitol, and (d) D-iduronate.

3. Describe the general structural features of glycosaminoglycans.

4. Honey is an emulsion of microcrystalline D-fructose and D-
glucose. Although D-fructose in polysaccharides exists mainly in
the furanose form, solution or crystalline D-fructose (as in honey)
is a mixture of several forms with /3-D-fructopyranose (67%) and
/3-D-fructofuranose (25%) predominating. Draw the Fischer pro-
jection for D-fructose and show how it can cyclize to form both of
the cyclized forms above.

5. Sialic acid (W-acetyl-a-D-neuraminic acid) is often found in
TV-linked oligosaccharides that are involved in cell-cell interactions.
Cancer cells synthesize much greater amounts of sialic acid than
normal cells. Derivatives of sialic acid have been proposed as anti-
cancer agents to block cell-surface interactions between normal
and cancerous cells. Answer the following questions about the
structure of sialic acid.

(a) Is it an a or a P anomeric form?

(b) Will sialic acid mutorotate between a and p anomeric forms?

(d) Will the open-chain form of sialic acid be an aldehyde or a
ketone?

(e) How many chiral carbons are there in the sugar ring?

Sialic acid

6. How many stereoisomers are possible for glucopyranose and for
fructofuranose? How many are D sugars in each case, and how
many are L sugars?

7. Draw the structure of each of the following molecules and label
each chiral carbon with an asterisk:

(a) cr-D-Glucose 1 -phosphate.

(b) 2-Deoxy-/3-D-ribose 5-phosphate.

(d) L-Glucuronate.

8. In aqueous solution, almost all D-glucose molecules (>99%) are
in the pyranose form. Other aldoses have a greater proportion of
molecules in the open-chain form. D-Glucose may have evolved
to be the predominant hexose because it is less likely than its iso-
mers to react with and damage cellular proteins. Explain why D-
glucose reacts less than other aldoses with the amino groups of
proteins.

9. Why is the /3-D-glucopyranose form of glucose more abundant
than a-D-glucopyranose in aqueous solution?

10. The relative orientations of substituents on ribose rings are deter-
mined by the conformation of the ring itself. If the ribose is part
of a polymeric molecule, then ring conformation will affect over-
all polymer structure. For example, the orientation of ribose
phosphate substituents connecting monomeric nucleoside units
is important in determining the overall structure of nucleic acid
molecules. In one major form of DNA (B-DNA), the ribofura-
nose rings adopt an envelope conformation in which C-2' carbon
is above the plane defined by C-l, C-3, C-4, and the ring oxygen
(C-2' endo conformation). Draw the envelope structure of D-
ribose 5-phosphate with a nucleoside base (B) attached in a
P -anomeric position at the C-l carbon.

11. In a procedure for testing blood glucose, a drop of blood is placed
on a paper strip impregnated with the enzyme glucose oxidase
and all the reagents necessary for the reaction

(3-D-G lucose + 0 2 » D-Gluconolactone + H 2 0 2

The H 2 0 2 produced causes a color change on the paper that indi-
cates how much glucose is present. Since glucose oxidase is spe-
cific for the p anomer of glucose, why can the total blood glucose
be measured?

12. Sucralose (registered under the brand name Splenda®) is a non-
nutritive (noncaloric) sweetener that is approximately 600 times
sweeter than sugar. Since sucralose is heat stable, it can be used in
cooking and baking. The structure of sucralose is shown below.

254 CHAPTER 8 Carbohydrates

Name the disaccharide that is used as a starting substrate for the
synthesis of sucralose. What chemical modifications have been
made to the starting disaccharide?

13 . Draw Haworth projections for the following glycosides:

(a) Isomaltose [ct-D-glucopyranosyl- ( 1 — » 6 ) -a-D-glucopyranose] .

(b) Amygdalin, a compound in the pits of certain fruits, which
has a — CH(CN)C 6 H 5 group attached to C-l of /3 -d-
glucopyranosyl-(l — > 6)-/3-D-glucopyranose.

14 . Keratan sulfate is a glycosaminoglycan composed primarily of the
following repeating disaccharide unit: — Gal /3(1 — » 4) GlcNAc6S
/3( 1 — » 3) — . The acetylated sugar has a sulfate ester on C-6. Ker-
atan sulfate is found in cornea, bone, and cartilage aggregated
with other glycosaminoglycans such as chondroitin sulfate. Draw
a Haworth projection of the repeating disaccharide unit found in
keratan sulfate.

15 . A number of diseases result from hereditary deficiencies in spe-
cific glycosidases. In these diseases, certain glycoproteins are in-
completely degraded and oligosaccharides accumulate in tissues.
Which of the XT-linked oligosaccharides in Figure 8.35 would be
affected by deficiencies of the following enzymes?

(a) iV-Acetyl-/3-glucosaminyl asparagine amidase

(b) /3-Galactosidase

(d) Fucosidase

16 . A carbohydrate-amino acid polymer that is a potent inhibitor of
influenza virus has been synthesized. The virus is thought to be
inactivated when multiple sialyl groups bind to viral surface pro-
teins. Draw the chemical structure of the carbohydrate portion of
this polymer (below, where X represents the rest of the polymer).

NeuNAc a -( 2 -> 3) Gal /3-(l -> 4) Glu /3-(l -> )-X

17 . Imagine that you could take a pill containing /3-glucosidase. If,
after taking this pill, you ate this textbook, what would it taste
like? Would it taste any different if you could marinate it
overnight in a solution containing /3-glucosidase? Should pub-
lishers use flavored ink in order to encourage students to eat their
textbooks?

Selected Readings

General

Collins, R M., ed. (1987). Carbohydrates (London
and New York: Chapman and Hall).

El Khadem, H. S. (1988). Carbohydrate Chemistry:
Monosaccharides and Their Derivatives (Orlando,
FL: Academic Press).

Li, X., Glaser, D., Li, W., Johnson, W. E., O’Brien,

S. J., Beauchamp, G. K., and Brand, J. G. (2009).
Analyses of sweet receptor gene (Taslr2) and pref-
erence for sweet stimuli in species of Carnivora.

/. Hered. 100 (Supplement 1):S90-S100.

Li, X., Li, W., Wang, H., Cao, J., Maehashi, K.,
Huang, L., Bachmanov, A. A., Reed, D. R.,
Legrand-Defretin, V., Beauchamp, G. K., and
Brand, J. G. (2005). Pseudogenization of a sweet-
receptor gene accounts for cats’ indifference to-
ward sugar. PloS Genet. 1(1): e3. DOL10.1371/
journal.pgen.0010003

Nodulation Factors

Denarie, J., and Debelle, L. (1996). Rhizobium
lipo-chitooligosaccharide nodulation factors:
signaling molecules mediating recognition
and morphogenesis. Annu. Rev. Biochem.
65:503-535.

Madsen, L. H., Tirichine, L., Jurkiewicz, A., Sullivan,
J. T., Heckmann, A. B., Bek, A. S., Ronson, C. W.,
James, E. K., and Stougaard, J. (2010). The molec-
ular network governing nodule organogenesis and
infection in the model legume Lotus jap onicus.
Nature Communications.

DOI: 10. 1038/ncommsl009

Mergaert, P., Van Montagu, M., and Holsters, M.
(1997). Molecular mechanisms of Nod factor
diversity. Mol. Microbiol. 25:81 1-817.

Thoden, J. B., Kim, J., Raushel, L. M., and
Holden, H. M. (2002). Structural and kinetic
studies of sugar binding to galactose mutarotase
from Lactococcus lactis. J. Biol. Chem.
277:45458-45465.

Proteoglycans

Heinegard, D., and Oldberg, A. (1989). Structure
and biology of cartilage and bone matrix
noncollagenous macromolecules. FASEB J.
3:2042-2051.

Iozzo, R. V. (1999). The biology of the small
leucine-rich proteoglycans: functional network
of interactive proteins. /. Biol. Chem.
274:18843-18846.

Iozzo, R. V., and Murdoch, A. D. (1996). Proteo-
glycans of the extracellular environment: clues
from the gene and protein side offer novel per-
spectives in molecular diversity and function.
FASEB J. 10:598-614.

Kjellen, L., and Lindahl, U. (1991). Proteoglycans:
structures and interactions. Annu. Rev. Biochem.
60:443-475.

Whitfield, C. (2006) Biosynthesis and assembly of
capsular polysaccharides in Escherichia coli. Annu.
Rev. Biochem. 75:39-68.

Glycoproteins

Drickamer, K., and Taylor, M. E. (1998). Evolving
views of protein glycosylation. Trends Biochem.

Sci. 23:321-324.

Dwek, R. A., Edge, C. J., Harvey, D. J., Wormald,
M. R., and Parekh, R. B. (1993). Analysis of
glycoprotein-associated oligosaccharides.

Annu. Rev. Biochem. 62:65-100.

Fudge, D. S., Levy, N., Chiu, S., and Gosline,

J. M. (2005). Composition, morphology and
mechanics of hagfish slime. /. Exp. Biol.
208:4613-4625.

Selected Readings 255

Lairson, L. L., Henrissat, B., Davies, G., and With-
ers, S. G. (2008). Glycosyltransferases: structures,
functions, and mechanisms. Annu Rev Biochem.
77:5 21-555.

Lechner, J., and Wieland, F. (1989). Structure and
biosynthesis of prokaryotic glycoproteins. Annu.
Rev. Biochem. 58:173-194.

Marionneau, S., Caileau-Thomas, A., Rocher,

J., Le Moullac-Vaidye, B. Ruvoen, N., Clement, M.,
and Le Pendu, J. (2001). ABH and Lewis histo-

blood group antigens, a model for the meaning of
oligosaccharide diversity in the face of a changing
world. Biochimie. 83:565-573.

Patenaude, S. I., Seto, N. O. L., Borisova, S. N.,
Szpacenko, A., Marcus, S. L., Palcic, M. M., and
Evans, S. V. (2002). The structural basis for speci-
ficity in human ABO(H) blood group biosynthe-
sis. Nat. Struct. Biol. 9:685-690.

Rademacher, T. W., Parekh, R. B., and Dwek, R. A.
(1988). Glycobiology. Annu. Rev. Biochem.
57:785-838

Rudd, P. M., and Dwek, R. A. (1997). Glycosyla-
tion: heterogeneity and the 3D structure of pro-
teins. Crit. Rev. Biochem. Mol. Biol. 32:1-100.

Strous, G. J., and Dekker, J. (1992). Mucin-type
glycoproteins. Crit. Rev. Biochem. Mol. Biol.
27:57-92.

o c

_ o

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° o

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° c

o o

Lipids and Membranes

I n this chapter, we consider lipids, ( lipo -, fat) a third major class of biomolecules.
Lipids — like proteins and carbohydrates — are essential components of all living
organisms. However, unlike these other types of biomolecules, lipids have widely
varied structures. They are often defined as water- insoluble (or only sparingly soluble)
organic compounds found in biological systems but that’s a very broad definition.
Lipids are very soluble in nonpolar organic solvents. They are either hydrophobic (non-
polar) or amphipathic (containing both nonpolar and polar regions).

We begin this chapter with a discussion of the structures and functions of the dif-
ferent classes of lipids. In the second part of the chapter, we examine the structures of
biological membranes whose properties as cellular barriers depend on the properties of
their lipids. Finally, we describe the principles of membrane transport and transmem-
brane signaling pathways.

In this article , we therefore present
and discuss a fluid mosaic model of
membrane structure ; and propose
that it is applicable to most biologi-
cal membranes ; such as plasmalem-
mal and intracellular membranes ,
including the membranes of different
cell organelles such as mitochondria
and chloroplasts .

— S.J. Singer and
G.L. Nicholson (1972)

9.1 Structural and Functional Diversity
of Lipids

Figure 9.1 shows the major types of lipids and their structural relationships to one
another. The simplest lipids are the fatty acids that have the general formula R —
COOH, where R represents a hydrocarbon chain composed of various lengths of
— CH 2 - (methylene) units. Fatty acids are components of many more complex types of
lipids, including triacylglycerols, glycerophospholipids, and sphingolipids. Lipids con-
taining phosphate groups are called phospholipids and lipids containing both sphingo-
sine and carbohydrate groups are called glycosphingolipids. Steroids, lipid vitamins, and
terpenes are related to the five-carbon molecule isoprene and are therefore called
isoprenoids. The name terpenes has been applied to all isoprenoids but usually is re-
stricted to those that occur in plants.

Lipids have diverse biological functions as well as diverse structures. Biological
membranes contain a variety of amphipathic lipids including glycerophospholipids and

Top: Ribbon structure of the transmembrane portion of porin FhuA from Escherichia coli (see Figure 9.28).

256

9.2 Fatty Acids 257

LIPIDS

Fatty acids

Eicosanoids

Triacylglycerols Waxes Sphingolipids

Glycerophospholipids

Ceramides

Plasmalogens Phosphatidates Sphingomyelins

Phosphatidyl- Phosphatidyl- Phosphatidyl- Phosphatidyl- Other

ethanolamines serines cholines inositols phospholipids

Phospholipids

sphingolipids. In some organisms, triacylglycerols (fats and oils) function as intracellu-
lar storage molecules for metabolic energy. Fats also provide animals with thermal insu-
lation and padding. Waxes in cell walls, exoskeletons, and skins protect the surfaces of
some organisms. Some lipids have highly specialized functions. For example, steroid
hormones regulate and integrate a host of metabolic activities in animals and
eicosanoids participate in the regulation of blood pressure, body temperature, and
smooth-muscle contraction in mammals. Gangliosides and other glycosphingolipids
are located at the cell surface and can participate in cellular recognition.

9.2 Fatty Acids

More than 100 different fatty acids have been identified in various species. Fatty
acids differ from one another in the length of their hydrocarbon tails, the number of
carbon-carbon double bonds, the positions of the double bonds in the chains, and the
number of branches. Some of the fatty acids commonly found in mammals are shown
in Table 9.1.

All fatty acids have a carboxyl group ( — CO OH) at their “head ” This is why they
are acids. The p K a of this group is about 4.5 to 5.0 so it is ionized at physiological pH
( — COO - ). Fatty acids are a form of detergent because they have a long hydrophobic
tail and a polar head (Section 2.4). As expected, the concentration of free fatty acid in
cells is quite low because high concentrations of free fatty acids could disrupt mem-
branes. Most fatty acids are components of more complex lipids. They are joined to
other molecules by an ester linkage at the terminal carboxyl group.

Fatty acids can be referred to by either International Union of Pure and Applied
Chemistry (IUPAC) names or common names. Common names are used for the most
frequently encountered fatty acids.

The number of carbon atoms in the most abundant fatty acids ranges from 12 to 20
and is almost always an even number since fatty acids are synthesized by the sequen-
tial addition of two-carbon units. In IUPAC nomenclature, the carboxyl carbon is la-
beled C-l and the remaining carbon atoms are numbered sequentially. In common

Steroids Lipid Terpenes
vitamins

Isoprenoids

Cerebrosides

Gangliosides

Other

glycosphingolipids

Glycosphingolipids

▲ Figure 9.1

Structural relationships of the major classes
of lipids. Fatty acids are the simplest lipids.
Many other types of lipids either contain or
are derived from fatty acids. Glycerophospho-
lipids and sphingomyelins contain phosphate
and are classified as phospholipids. Cerebro-
sides and gangliosides contain sphingosine
and carbohydrate and are classified as gly-
cosphingolipids. Steroids, lipid vitamins,
and terpenes are called isoprenoids because
they are related to the five-carbon molecule
isoprene rather than to fatty acids.

Fatty acid biosynthesis is discussed in
Chapter 16.

258 CHAPTER 9 Lipids and Membranes

Table 9.1 Some common fatty acids (anionic forms)

Number of
carbons

Number of
double bonds

Common

name

IUPAC name

Molecular formula

Melting
point, °C

Laurate

Dodecanoate

CH 3 (CH 2 ) 10 COO©

Myristate

Tetradecanoate

CH 3 1(CH 2 1) 12 COO e

Palmitate

Hexadecanoate

CH 3 1(CH 2 1) 14 COO e

Stearate

Octadecanoate

CH 3 (CH 2 ) 16 COO©

Arachidate

Eicosanoate

CH 3 (CH 2 ) 18 COO©

Behenate

Docosanoate

CH 3 (CH 2 ) 20 COO e

Lignocerate

Tetracosanoate

CH 3 (CH 2 ) 22 COO e

Palmitoleate

cis- A 9 -Hexadecenoate

CH 3 (CH 2 ) 5 CH = CH(CH 2 ) 7 COO©

-0.5

Oleate

c/s- A 9 -Octadecenoate

CH 3 (CH 2 ) 7 CH = CH(CH 2 ) 7 COO e

Linoleate

cis, c/s- A 9,1 2 -Octadecadienoate

CH 3 (CH 2 ) 4 (CH = CHCH 2 ) 2 (CH 2 ) 6 COO e

-9

Linolenate

all c/s- A 9/1 2/1 5 -Octadecatrienoate

CH 3 CH 2 (CH = CHCH 2 ) 3 (CH 2 ) 6 COO e

-17

Arachidonate

all c/s- A 5,8,1 1 ^ 4 -Eicosatetraenoate

CH 3 (CH 2 ) 4 (CH = CHCH 2 ) 4 (CH 2 ) 2 COO®

-49

nomenclature, Greek letters are used to identify the carbon atoms. The carbon adjacent
to the carboxyl carbon (C-2 in IUPAC nomenclature) is designated a, and the other
carbons are lettered /?, y, <5, and £ and so on (Figure 9.2). The Greek letter co (omega)
specifies the carbon atom farthest from the carboxyl group, whatever the length of the
hydrocarbon tail, (co is the last letter in the Greek alphabet.)

Fatty acids without a carbon-carbon double bond are classified as saturated,
whereas those with at least one carbon-carbon double bond are classified as unsaturated.
Unsaturated fatty acids with only one carbon-carbon double bond are called
monounsaturated and those with two or more are called polyunsaturated. The configuration
of the double bonds in unsaturated fatty acids can be either cis or trans . The configura-
tion is usually cis in naturally occurring fatty acids (see Box. 9.2).

The positions of double bonds are indicated by the symbol A n in IUPAC nomen-
clature, where the superscript n indicates the lower-numbered carbon atom of each

BOX 9.1 COMMON NAMES OF THE FATTY ACIDS

Laurate

Myristate

Palmitate

Stearate

Arachidate

Behenate

Lignocerate

Oleate

Linoleate

present in oil from the laurel plant ( Laurus
nobilis) (1873)

oil from nutmeg (Myristica fragrans) (1848)
from palm oil (1857)

from French stearique referring to fat from
steers, or tallow (1831)

present in oil from peanuts ( Arachis hypogaea )
(1866)

a corruption of “ben” from ben- nut = seeds of

the Horseradish tree (1873)

probably from Latin lignum (“wood”) (-1900)

from Latin oleum (“oil”) (1899)

found in linseed oil ( lin + oleate ) (1857)

▲ The African oil palm tree, Elaeis guineensis. Palm oil is a complex
mixture of saturated and unsaturated fatty acids but palmitate
makes up 44% of the total. The presence of such a large amount of
saturated fatty acid means that palm oil is a semisolid at room tem-
perature. It can never be “virgin” or “extra virgin” (see Box 16.6).

9.2 Fatty Acids 259

o©

0= c

’\

a CH

/ 2

/jCH

3 \

rCH 2

/ 4

5 ch 2

5 \

«CH

7 \

/ 8

9 \

/i°

« CH

Fatty

acid

Fatty

acyl

group

Hydrocarbon

tail

◄ Figure 9.2

Structure and nomenclature of fatty acids. Fatty
acids consist of a long hydrocarbon tail ter-
minating with a carboxyl group. Since the
p K a of the carboxyl group is approximately
4.5 to 5.0, fatty acids are anionic at physio-
logical pH. In IUPAC nomenclature, carbons
are numbered beginning with the carboxyl
carbon. In common nomenclature, the car-
bon atom adjacent to the carboxyl carbon is
designated a, and the remaining carbons
are lettered p, y S, and so on. The carbon
atom farthest from the carboxyl carbon is
designated the co carbon, whatever the length
of the tail. The fatty acid shown, laurate
(or dodecanoate), has 12 carbon atoms and
contains no carbon-carbon double bonds.

double-bonded pair (Table 9.1). The double bonds of most polyunsaturated fatty acids
are separated by a methylene group and are therefore not conjugated.

A shorthand notation for identifying fatty acids uses two numbers separated by
a colon — the first refers to the number of carbon atoms in the fatty acid and the second
refers to the number of carbon-carbon double bonds, with their positions indicated
as superscripts following a Greek symbol, A. In this notation, palmitate is written as
16:0, oleate as 18:1 A 9 , and arachidonate as 20:4 A 5,8,11,14 . Unsaturated fatty acids can

BOX 9.2 TRANS FATTY ACIDS AND MARGARINE

The configuration of most double bonds in unsaturated fatty
acids is cis but some fatty acids in the human diet have the
trans configuration. Trans fatty acids can come from animal
sources such as dairy products and ruminant meats. However,
most of the edible trans fatty acids consumed in Western in-
dustrialized countries are present as hydrogenated vegetable
oils in some margarines or shortenings. Dietary trans
monounsaturated fatty acids can increase plasma levels of
cholesterol and triglycerides and their ingestion may increase
the risk of cardiovascular disease. More work is required to
establish the exact level of risk.

Plant oils such as corn oil and sunflower oil can be converted
to “spreadable” semisolid substances known as margarines.
Margarines can be produced by the partial or complete hy-
drogenation of double bonds in plant oils. The hydrogenation
process itself not only saturates the carbon-carbon double
bonds of fatty acid esters but can also change the configuration
of the remaining double bonds from cis to trans. The physical
properties of these trans fatty acids are similar to those of sat-
urated fatty acids.

In order to reduce consumption of trans fatty acids,
many margarines are now produced from plant oils without
hydrogenation by adding other edible components such as
skim milk powder.

COOH

▲ Cis and transforms of A 9 -octadecanoate. (Left) Oleate (c/'s- A 9 -octade-
canoate). (Right) the trans configuration after hydrogenation.

260

CHAPTER 9 Lipids and Membranes

t Figure 9.3

Structures of three C 18 fatty acids, (a) Stearate
(octadecanoate), a saturated fatty acid, (b)
Oleate (c/s- A 9 -octadecenoate) a monounsat-
urated fatty acid, (c) Linolenate (all-c/'s-
A 9 ,i2T5_ oc t ac j eca t r j enoa t e ) a polyunsaturated
fatty acid. The cis double bonds produce
kinks in the tails of the unsaturated fatty
acids. Linolenate is a very flexible molecule
that can assume a variety of conformations.

also be described by the location of the last double bond in the chain. This double bond
is usually found three, six, or nine carbon atoms from the end of the chain. Such fatty
acids are called co - 3 (e.g., 18:3 A 9,12,15 ), co - 6 (e.g., 18:2 A 9,12 ), or co - 9 (e.g., 18:1 A 9 ).

The physical properties of saturated and unsaturated fatty acids differ considerably.
Typically, saturated fatty acids are waxy solids at room temperature (22°C) whereas un-
saturated fatty acids are liquids at this temperature. The length of the hydrocarbon
chain of a fatty acid and its degree of unsaturation influence the melting point. Com-
pare the melting points listed in Table 9.1 for the saturated fatty acids laurate (12:0),
myristate (14:0), and palmitate (16:0). As the lengths of the hydrocarbon tails increase,
the melting points of the saturated fatty acids also increase. The number of van der
Waals interactions among neighboring hydrocarbon tails increases as the tails get longer
so more energy is required to disrupt the interactions.

Compare the structures of stearate (18:0), oleate (18:1), and linolenate (18:3) in
Figures 9.3 and 9.4. The saturated hydrocarbon tail of stearate is flexible since rotation
can occur around every carbon-carbon bond. In a crystal of stearic acid, the hydrocar-
bon chains are extended and pack together closely. The presence of cis double bonds in
oleate and linolenate produces pronounced bends in the hydrocarbon chains since rota-
tion around double bonds is hindered. These bends prevent close packing and extensive
van der Waals interactions among the hydrocarbon chains. Consequently, cis unsatu-
rated fatty acids have lower melting points than saturated fatty acids. As the degree of
unsaturation increases, fatty acids become more fluid. Note that stearic acid (melting
point 70°C) is a solid at body temperature but oleic acid (melting point 13°C) and
linolenic acid (melting point — 17°C) are both liquids.

As mentioned earlier, free fatty acids occur only in trace amounts in living cells.
Most fatty acids are esterified to glycerol or other backbone compounds to form more
complex lipid molecules. In esters and other derivatives of carboxylic acids, the RC = O
moiety contributed by the acid is called the acyl group. In common nomenclature,

(a)

0 G

= c

2CH 2

3CH 2

4CH 2

5CH 2

6CH 2

7CH 2

8CH 2

9CH 2

10CH 2

"CH 2

12CH 2

'3CH 2

14CH 2

'5CH 2

16CH 2

17CH 2

18CH 3

Stearate

(b)

0 G

0 = C

2CH 2

3CH 2

4CH 2

3CH 2

6CH 2

7CH 2

8CH 2

(c)

H— 9 C

^ 11

10 C— ch 2

/ \ 13

H H 2 C— CH 2

12 \ z

\ 15

h 2 c — ch 2

^ 14 \

\ 17

h 2 c— ch 2

z i6 y z

18CH3

0 G

0= c

2CH 2

3CH 2

4CH 2

5CH 2

6CH 2

7CH 2

.ch 2

1 oC=C 9

/ \

H H

;ch 2

15C-

17 16 //

h 2 c — c

/ \

l-UO

Oleate

Linolenate

9.3 Triacylglycerols 261

◄ Figure 9.4

Stearate (left), oleate (center), and linolenate
(right). Color key: carbon, grey; hydrogen,
white; oxygen, red.

(a)

(b)

1 2 I 3

h 2 c — c — ch 2

1 I I

OH OH OH
H 2 C — ch — ch 2

complex lipids that contain specific fatty acyl groups are named after the parent fatty
acid. For example, esters of the fatty acid laurate are called lauroyl esters, and esters of
linoleate are called linoleoyl esters. (A lauryl group is the alcohol analog of the lauroyl
acyl group). The relative abundance of particular fatty acids varies with the type of or-
ganism, type of organ (in multicellular organisms), and food source. The most abundant
fatty acids in animals are usually oleate (18:1), palmitate (16:0), and stearate (18:0).

Mammals require certain dietary polyunsaturated fatty acids that they cannot syn-
thesize, such as linoleate (18:2 A 9,12 ) and linolenate (18:3 A 9,12,15 ). These fatty acids are
called essential fatty acids. Mammals can synthesize other polyunsaturated fatty acids
from an adequate supply of linoleate and linolenate. (Recall that many vitamins are also
essential components of the mammalian diet because mammals cannot synthesize
them. In addition to vitamins and essential fatty acids, we will see in Chapter 17 that
many amino acids cannot be synthesized in mammals.)

Linolenate is an omega- 3 (ft) - 3) fatty acid since the last double bond is three carbon
atoms from the tail end of the molecule. Omega-3 fatty acids are very popular dietary
supplements. They are enriched in fish oils, which is why many people recommend that
you include fish and fish oils in your diet. Linolenate is an essential fatty acid so your diet
must include an adequate supply of this omega-3 fatty acid. This adequate amount is
readily supplied in the typical diet of people all over the world, which is why essential
fatty acid deficiency is rare. The market for supplemental omega-3 fatty acids is driven by
other factors. The most important benefit is protection against cardiovascular disease.
The scientific evidence indicates that extra amounts of omega- 3 fatty acids provide a
small benefit in terms of reducing the risk of heart attacks, particularly a second heart at-
tack. None of the other claims are based on reproducible double-blind test results after
controlling for other factors. Eating fish, for example, will not make you smarter.

Many fatty acids besides those listed in Table 9.1 are present in nature. For example,
fatty acids containing cyclopropane rings are found in bacteria. Branched-chain fatty
acids are common components of bacterial membranes and also occur on the feathers
of ducks. Many other fatty acids are rare and have highly specialized functions.

9.3 Triacylglycerols

As their name implies, triacylglycerols (historically called triglycerides) are composed of
three fatty acyl residues esterified to glycerol, a three-carbon sugar alcohol (Figure 9.5).
Triacylglycerols are very hydrophobic.

▲ Figure 9.5

Structure of a triacylglycerol. Glycerol (a) is
the backbone to which three fatty acyl residues
are esterified (b). Although glycerol is not
chiral, C-2 of a triacylglycerol is chiral when
the acyl groups bound to C-l and C-3 (Ri and
R 3 ) differ. The general structure of a triacyl-
glycerol is shown in (c), oriented for compar-
ison with the structure of L-glyceraldehyde
(Figure 8.1). This orientation allows stere-
ospecific numbering of glycerol derivatives
with C-l at the top and C-3 at the bottom.

262 CHAPTER 9 Lipids and Membranes

▲ Figure 9.6

Adipocytes. This is a colorized scanning
electron micrograph of clusters of adipocytes.
A fat droplet occupies most of the volume of
each adipocyte.

The structures and functions of lipopro-
teins are discussed in Section 16.1B.

KEY CONCEPT

Glycerophospholipids have polar heads
and long, hydrophobic fatty acid tails.

KEY CONCEPT

Many important lipids are derivatives of
glycerol (see Box 16.1).

▲ Yellow jacket wasp. The venom of wasps,
bees, and snakes contains phospholipases.

Fats and oils are mixtures of triacylglycerols. They can be solids (fats) or liquids
(oils), depending on their fatty acid compositions and on the temperature. Triacylglyc-
erols containing only saturated long chain fatty acyl groups tend to be solids at body
temperature and those containing unsaturated or short chain fatty acyl groups tend to
be liquids. A sample of naturally occurring triacylglycerols can contain as many as 20 to
30 molecular species that differ in their fatty acid constituents. Tripalmitin, found in
animal fat, contains three residues of palmitic acid. Triolein, which contains three oleic
acid residues, is the principal triacylglycerol in olive oil.

In most cells, triacylglycerols coalesce as fat droplets. These droplets are sometimes
seen near mitochondria in cells that rely on fatty acids for metabolic energy. In mam-
mals, most fat is stored in adipose tissue that is composed of specialized cells known as
adipocytes. Each adipocyte contains a large fat droplet that accounts for nearly the en-
tire volume of the cell (Figure 9.6). Although distributed throughout the bodies of
mammals, most adipose tissue occurs just under the skin and in the abdominal cavity.
Extensive subcutaneous fat serves both as a storage depot for energy and as thermal in-
sulation and is especially pronounced in aquatic mammals.

9.4 Glycerophospholipids

Triacylglycerols are not found in biological membranes. The most abundant lipids in
most membranes are glycerophospholipids (also called phosphoglycerides). Glyc-
erophospholipids, like triacylglycerols, have a glycerol backbone. The simplest glyc-
erophospholipids are the, phosphatidates — they consist of two fatty acyl groups
esterified to C-l and C-2 of glycerol 3-phosphate (Table 9.2). Note that there are three
fatty acyl groups esterified to glycerol in triacylglycerols whereas there are only two fatty
acyl groups (R x and R 2 ) in the glycerophospholipids. The distinguishing feature of
the glycerophospholipids is the presence of a phosphate group on C-3 of the glycerol
backbone. The structures of glycerophospholipids can be drawn as derivatives of
L-glycerol 3-phosphate with the C-2 substituent on the left in a Fischer projection, as in
Table 9.2. For simplicity, we usually show these compounds as stereochemically uncom-
mitted structures.

Phosphatidates are present in small amounts as intermediates in the biosynthe-
sis or breakdown of more complex glycerophospholipids. In most glycerophospho-
lipids, the phosphate group is esterified to both glycerol and another compound
bearing an — OH group. Table 9.2 lists some common types of glycerophospho-
lipids. Note that glycerophospholipids are amphipathic molecules with a polar head
and long, nonpolar tails. The structures of three types of glycerophospholipids —
phosphatidylethanolamine, phosphatidylserine, and phosphatidylcholine — are shown
in Figure 9.7.

Each type of glycerophospholipid consists of a family of molecules with the same
polar head group and different fatty acyl chains. For example, human red blood cell
membranes contain at least 21 different species of phosphatidylcholine that differ from
one another in the fatty acyl chains esterified at C-l and C-2 of the glycerol backbone.
In general, glycerophospholipids have saturated fatty acids esterified to C-l and unsatu-
rated fatty acids esterified to C-2. The major membrane glycerophospholipids in
Escherichia coli are phosphatidylethanolamine and phosphatidylglycerol.

A variety of phospholipases can be used to dissect glycerophospholipid structures
and determine the identities of their individual fatty acids. The specific positions of
fatty acids in glycerophospholipids can be determined by using phospholipase A x and
phospholipase A 2 that specifically catalyze the hydrolysis of the ester bonds at C- 1 and
C-2, respectively (Figure 9.8). Phospholipase A 2 is the major phospholipase in pancre-
atic juice and it is responsible for the digestion of membrane phospholipids in the diet.
It is also present in snake, bee, and wasp venom. High concentrations of the products of
the action of phospholipase A 2 can disrupt cell membranes. Thus, injection of snake
venom into the blood can result in life-threatening lysis of the membranes of red blood
cells. Phospholipase C catalyzes hydrolysis of the P — O bond between glycerol and

9.5 Sphingolipids 263

Table 9.2 Some common types of glycerophospholipids

(Ri)

c— o— ch 2

X = rest of polar head

(R 2 )

o — X

Precursor of X
(HO — X)

Formulas of — O

-X

Name of resulting
glycerophospholipid

Water

— H

Phosphatidate

Choline

— CH 2 CH 2 N(CH 3 ) 3

Phosphatidylcholine

Ethanolamine

— ch 2 ch 2 nh 3

Phosphatidylethanolamine

Serine

NhU

— CH 2 — CH

\oo 0

Phosphatidylserine

Glycerol

— ch 2 ch — ch 2 oh

Phosphatidylglycerol

0 CH 2 OCR 3

r 4 coch

Phosphatidyl-

glycerol

— ch 2 ch — ch 2 — 0

— p-

o e

-o— ch 2

Diphosphatidylglycerol

(Cardiolipin)

myo- Inositol

H OH

1/ DH H\j

1 |C OH HO A

Phosphatidyl inositol

H 0H
H H

phosphate to liberate diacylglycerol. Phospholipase D converts glycerophospholipids to
phosphatidates.

Plasma logens are the other major type of glycerophospholipids. They differ from phos-
phatidates because the hydrocarbon substituent on the C- 1 hydroxyl group of glycerol
is attached by a vinyl ether linkage rather than an ester linkage (Figure 9.9). Ethanolamine
or choline is commonly esterified to the phosphate group of plasmalogens. Plasmalogens
account for about 23% of the glycerophospholipids in the human central nervous system
and are also found in the membranes of peripheral nerve and muscle tissue.

9.5 Sphingolipids

Sphingolipids are the second most abundant lipids in plant and animal membranes. In
mammals, sphingolipids are particularly abundant in tissues of the central nervous sys-
tem. Most bacteria do not have sphingolipids. The structural backbone of sphingolipids
is sphingosine (trans- 4-sphingenine), an unbranched C 18 alcohol with a trans double

264 CHAPTER 9 Lipids and Membranes

(R,) (R 2 )

Phosphatidylethanolamine

Phosphatidylserine

(Ri) (R 2 )

Phosphatidylcholine

^ Polar heads
(hydrophilic)

^ Nonpolar tails
(hydrophobic)

▲ Figure 9.7

Structures of (a) phosphatidylethanolamine,
(b) phosphatidylserine, and (c) phosphatidyl-
choline. Functional groups derived from es-
terified alcohols are shown in blue. Since
each of these lipids can contain many com-
binations of fatty acyl groups, the general
name refers to a family of compounds, not
to a single molecule.

bond between C-4 and C-5, an amino group at C-2, and hydroxyl groups at C-l and C-3
(Figure 9.10a). Ceramide consists of a fatty acyl group linked to the C-2 amino group of
sphingosine by an amide bond (Figure 9.10b). Ceramides are the metabolic precursors
of all sphingolipids. The three major families of sphingolipids are the sphingomyelins,
the cerebrosides, and the gangliosides. Of these, only sphingomyelins contain phosphate
and are classified as phospholipids; cerebrosides and gangliosides contain carbohydrate
residues and are classified as glycosphingolipids (Figure 9.1).

In sphingomyelins, phosphocholine is attached to the C-l hydroxyl group of a ce-
ramide (Figure 9.10c). Note the resemblance of sphingomyelin to phosphatidylcholine
(Figure 9.7c) — both molecules are zwitterions containing choline, phosphate, and two
long hydrophobic tails. Sphingomyelins are present in the plasma membranes of most
mammalian cells and are a major component of the myelin sheaths that surround cer-
tain nerve cells.

Figure 9.8 ►

Action of four phospholipases. Phospholipases
A 1? A 2 , C, and D can be used to dissect glyc-
erophospholipid structure. Phospholipases
catalyze the selective removal of fatty acids
from C-l or C-2 or convert glycerophospho-
lipids to diacylglycerols or phosphatidates.

H 2 C— CH— CH 2
O O

Phospholipase A ^ -| |- Phospholipase A 2

0 = c c = o

Phospholipase D

0 o— P = 0

I- Phospholipase C

Ri R 2 Rt

9.5 Sphingolipids 265

Cerebrosides are glycosphingolipids that contain one monosaccharide residue
attached by a /3-glycosidic linkage to C-l of a ceramide. Galactocerebrosides, also
known as galactosylceramides, have a single /3 -d - galactosyl residue as a polar head
group (Figure 9.11). Galactocerebrosides are abundant in nerve tissue and account for
about 15% of the lipids of myelin sheaths. Many other mammalian tissues contain glu-
cocerebrosides, ceramides with a /3-D-glucosyl head group. In some glycosphingolipids,
a linear chain of up to three more monosaccharide residues is attached to the galactosyl
or glucosyl moiety of a cerebroside.

Gangliosides are more complex glycosphingolipids in which oligosaccharide chains
containing N-acetylneuraminic acid (NeuNAc) are attached to a ceramide. NeuNAc
(Figure 8.15), an acetylated derivative of neuraminic acid, makes the head groups of
gangliosides anionic. The structure of a representative ganglioside, G M2 , is shown in
Figure 9.12. The M in G M2 stands for monosialo (i.e., one NeuNAc residue); G M2 was
the second monosialo ganglioside characterized, thus the subscript 2.

More than 60 varieties of gangliosides have been characterized. Their structural di-
versity results from variations in the composition and sequence of sugar residues. Gan-
glioside G M1 , for example, is similar to ganglioside G M2 shown in Figure 9.12 except
that it has an additional /3 -d- galactose residue attached to the terminal N-acetyl-/3-D-
galactosamine residue via a /3-( 1 —> 4) linkage. In all gangliosides, the ceramide is linked
through its C-l to a /3-glucosyl residue, which in turn is bound to a /3-galactosyl residue.

Gangliosides are present on cell surfaces with the two hydrocarbon chains of the
ceramide moiety embedded in the plasma membrane and the oligosaccharides on the

(a)

Sphingosine
(trans- 4-Sphingenine)

(c)

Sphingomyelin

CH q

©J

(b)

h 3 c — n — ch 3
oh 2
oh 2
o

Ceramide

0=P— 0°

©nh 3

ch 2

oh 2

0= P— o 0

Vinyl
ether -<
linkage

1 2 3 I

h 2 c — ch — ch 2

c = o

(Ri) (R 2 )

▲ Figure 9.9

Structure of an ethanolamine plasmalogen. A

hydrocarbon is linked to the C-l hydroxyl
group of glycerol to form a vinyl ether.

HO OH

I i 2 3 I

h 2 c — ch — ch

4CH

HC 5

/ CHl

C \ 2

/ CH 2

/CH2

ch 2

/ CH2

C h 2

/ CHi

C \ 2

/ c " 2

c „ 2

18CH 3

HO OH

I 1 2 3 I

H 2 c — CH — CH
NH

0 = c

(R)

4CH

HC 5

/ CH2

c x h 2

/ CH 2

c x h 2

/CH 2

c x h 2

/ CH 2

c x h 2

/CH 2

ch 2

18CH 3

H,c — CH — CH

o=c

(R)

4CH

HC 5

/ CH2

ch 2

/ CH 2

ch 2

/H2

ch 2

/ CH 2

ch 2

/ CH 2

ch 2

/ Hj

ch 2

i8 C H 3

Genetic defects associated with lipid
metabolism are described in Chapter 16.

◄ Figure 9.10

Structures of sphingosine, ceramide, and
sphingomyelin, (a) Sphingosine, the back-
bone for sphingolipids, is a long-chain alcohol
with an amino group at C-2. (b) Ceramides
have a long-chain fatty acyl group attached
to the amino group of sphingosine. (c) Sphin-
gomyelins have a phosphate group (red) at-
tached to the C-l hydroxyl group of a ceramide
and a choline group (blue) attached to the
phosphate.

266 CHAPTER 9 Lipids and Membranes

Ceramide

▲ Figure 9.1 1

Structure of a galactocerebroside. j8-D-Ga lactose
(blue) is attached to the C-l hydroxyl group of a
ceramide (black).

(a)

H sC x /H 2

A C \

h 2 c h

▲ Figure 9.13

Isoprene (2-methyl-1 ,3-butadiene), the basic
structural unit of isoprenoids. (a) Chemical
structure, (b) Carbon backbone, (c) Isoprene
unit where dashed lines represent covalent
bonds to a adjacent units.

I /3-D-Galactose

H 3 C /C ^0

A/-Acetyl-/3-D-galactosamine

extracellular surface. Gangliosides and other glycosphingolipids are part of the cell sur-
face repertoire of diverse oligosaccharide chains along with glycoproteins. Collectively,
these markers provide cells with distinguishing surface markers that can serve in cellular
recognition and cell-to-cell communication. Structures similar to the ABO blood group
antigens on the surface of human cells (Box. 8.3) can be oligosaccharide components of
glycosphingolipids in addition to being linked to proteins to form glycoproteins.

Genetically inherited defects in ganglioside metabolism are responsible for a
number of debilitating and often lethal diseases, such as Tay-Sachs disease and gener-
alized gangliosidosis. Certain rare genetic defects lead to deficiencies of enzymes re-
sponsible for the degradation of sphingolipids in the lysosomes of cells. In Tay-Sachs
disease, there is a deficiency of a hydrolase that catalyzes removal of N- acetylgalac-
tosamine from G M2 . Accumulation of G M2 causes lysosomes to swell leading to tissue
enlargement. In the central nervous tissue, where there is little room for expansion,
nerve cells die causing blindness, mental retardation, and death.

The exposed carbohydrates on the cell surface also provide convenient receptors
for bacteria, viruses, and toxins. For example, cholera toxin, produced by the bac-
terium Vibrio cholerae , binds to the ganglioside G M1 of intestinal epithelial cells. Bind-
ing stimulates entry of the toxin into the cells where it interferes with normal signaling
pathways leading to massive efflux of fluid into the intestine. This often produces
death by dehydration.

9.6 Steroids

Steroids are a third class of lipids found in the membranes of eukaryotes and, very rarely,
in bacteria. Steroids, along with lipid vitamins and terpenes, are classified as isoprenoids
because their structures are related to the five-carbon molecule isoprene (Figure 9.13).
Steroids contain four fused rings: three six- carbon rings designated A, B, and C and a five-
carbon D ring. The characteristic ring structure is derived from squalene (Figure 9.14a).
Substituents of the nearly planar ring system can point either down (the a configuration)
or up (the /3 configuration). The structures of several steroids are shown in Figure 9.14.

The steroid cholesterol is an important component of animal plasma membranes
but is less common in plants and absent from prokaryotes, protists, and fungi. These
species have other steroids (e.g., stigmasterol, ergosterol) that are very similar to choles-
terol. Cholesterol is actually a sterol because it has a hydroxyl group at C-3. Other
steroids include the sterols of plants, fungi, and yeast (which also have a hydroxyl group
at C-3); mammalian steroid hormones (such as estrogens, androgens, progestins, and

9.6 Steroids 267

◄ Figure 9.14

Structures of several steroids. Squalene (a) is
the precursor of most steroids. Steroids
contain four fused rings (lettered A, B, C,
and D). (b) Cholesterol, (c) Stigmasterol, a
common component of plant membranes.

(d) Testosterone, a steroid hormone involved
in male development in animals, (e) Sodium
cholate, a bile salt, which aids in the diges-
tion of lipids, (f) Ergosterol, a compound
from fungi and yeast.

Stigmasterol Testosterone

(a plant sterol) (a steroid hormone)

Sodium cholate Ergosterol

(a bile salt) (a sterol from fungi and yeast)

adrenal corticosteroids); and bile salts. These steroids differ in the length of the side chain
attached to C-17 and in the number and placement of methyl groups, double bonds, hy-
droxyl groups, and in some cases, keto groups. Prokaryotes use squalene and some re-
lated nonsteroid lipids that do not have the complete ring structure of the steroids.

Cholesterol plays an essential role in mammalian biochemistry. It is not only a
component of certain membranes but is also a precursor of steroid hormones and bile
salts. The fused ring system of cholesterol, shown from the side in Figure 9.15, makes it
less flexible than most other lipids. As a result, cholesterol modulates the fluidity of
mammalian cell membranes, as we will see later in this chapter.

Steroids are far more hydrophobic than glycerophospholipids and sphingolipids.
For example, free cholesteroPs maximal concentration in water is only 10 -8 M. Esterifi-
cation of a fatty acid to the C-3 hydroxyl group forms a cholesteryl ester (Figure 9.16).

▲ Figure 9.15

Cholesterol, (a) Bal l-and-stick model with
the oxygen atom (red) at the top. Hydrogen
atoms are not shown. The fused ring system
of cholestrol is almost planar, (b) Space-
filling model.

268 CHAPTER 9 Lipids and Membranes

Because the 3-acyl group of the ester is nonpolar, a cholesteryl ester is even more
hydrophobic than cholesterol itself. Cholesterol is converted to cholesteryl esters for
storage in cells or for transport through the bloodstream. Because they are essentially
insoluble in water, cholesterol and its esters must be complexed with phospholipids and
amphipathic proteins in lipoproteins for transport (Section 16. IB).

H 3 C - (CH 2 ) 14 — C - O - (CH 2 ) 29 CH 3

▲ Figure 9.17
Myricyl palmitate, a wax.

9.7 Other Biologically Important Lipids

There are many kinds of lipids not found in membranes. These include diverse com-
pounds such as waxes, eicosanoids, and some isoprenoids. Non-membrane lipids have a
variety of specialized functions — some of which we have already encountered (e.g.,
lipid vitamins).

Waxes are nonpolar esters of long-chain fatty acids and long chain monohydrox-
ylic alcohols. For example, myricyl palmitate, a major component of beeswax, is the
ester of palmitate (16:0) and the 30-carbon myricyl alcohol (Figure 9.17). The hy-
drophobicity of myricyl palmitate makes beeswax very insoluble and its high melt-
ing point (due to the long, saturated hydrocarbon chains) makes beeswax hard and
solid at typical outdoor temperatures. Waxes are widely distributed in nature. They
provide protective waterproof coatings on the leaves and fruits of certain plants and
on animal skin, fur, feathers, and exoskeletons. Ear wax, also known as cerumen
(from the Latin word cera , “wax”), is secreted by cells lining the auditory canal. It
serves to lubricate the canal and trap particles that could damage the eardrum. Ear
wax is a complex mixture made up mostly of long chain fatty acids, cholesterol, and
ceramides. It also contains squalene, triacylglycerols, and true waxes (about 10% of
the weight).

Eicosanoids are oxygenated derivatives of C 2 o polyunsaturated fatty acids such as
arachidonic acid. Some examples of eicosanoids are shown in Figure 9.18. Eicosanoids
participate in a variety of physiological processes and can also mediate many potentially
pathological responses. Prostaglandins are eicosanoids that have a cyclopentane ring.

▲ Earwax and beeswax are two examples of
naturally occurring waxes.

(b)

Prostaglandin E 2

▲ Figure 9.18

Structures of arachidonic acid (a) and three eicosanoids derived from it. Arachidonate is a C 2 o polyun-
saturated fatty acid with four c/'s double bonds.

9.8 Biological Membranes 269

(c)

(d)

Prostaglandin E 2 can cause constriction of blood vessels, and thromboxane A 2 is in- (a)
volved in the formation of blood clots that in some cases can block the flow of
blood to the heart or brain. Leukotriene D 4 , a mediator of smooth-muscle con-
traction, also provokes the bronchial constriction seen in asthmatics. Aspirin
(acetylsalicylic acid) alleviates pain, fever, swelling, and inflammation by inhibiting
the synthesis of prostaglandins (Box. 16.1).

Some nonmembrane lipids are related to isoprene (Figure 9.13) but they are (b)
not steroids. We encountered several of these lipids in Chapter 7. The lipid vitamins
A, E, and K are isoprenoids that contain long hydrocarbon chains or fused rings
(Section 7.14). Vitamin D is an isoprenoid derivative of cholesterol. There are sev-
eral carotenes related to retinol (vitamin A). The hydrophobic chain of ubiquinone
contains 6-10 isoprenoid units (Section 7.15).

Simple isoprenoids are often called terpenes. They have structures that reveal
their formation from isoprene units. Citral is a good example: it is present in many
plants and imparts a strong lemon odor (Figure 19.19a). Other isoprenoids are bac-
toprenol (undecaprenyl alcohol) (Figure 9.19b) and juvenile hormone I (Figure
9.19c) that regulates the expression of genes required for development in insects.
Isoprenoids similar to bactoprenol are important lipids in archaebacteria, where
they replace fatty acids in most membrane phospholipids (see Box 9.5).

Terpenes can be extensively modified to form a more complex class of lipid
called terpenoids. Many of these are cyclic compounds like limonene, which is re-
sponsible for the smell of oranges (Figure 19.19d). Gibberellins are multi-ring ter-
penoids that function as growth hormones in plants (Figure 19.19e).

9.8 Biological Membranes

Biological membranes define the external boundaries of cells and separate com-
partments within cells. They are essential components of all living cells. A typical
membrane consists of two layers of lipid molecules and many embedded proteins.

Biological membranes are not merely passive barriers to diffusion. They have a
wide variety of complex functions. Some membrane proteins serve as selective
pumps controlling the transport of ions and small molecules into and out of the
cell. Membranes are also responsible for generating and maintaining the proton
concentration gradients essential for the production of ATP. Receptors in mem-
branes recognize extracellular signals and communicate them to the cell interior.

Many cells have membranes with specialized structures. For example, many bac-
teria have double membranes: an outer membrane and an inner plasma membrane.
The liquid in the periplasmic space between these two membranes contains proteins
that carry specific solutes to transport proteins in the inner membrane. The solutes
then pass through the inner membrane by an ATP-dependent process. A mitochon-
drion’s smooth outer membrane has proteins that form aqueous channels while its
convoluted inner membrane is selectively permeable and has many membrane-
bound enzymes. The nucleus also has a double membrane — nuclear contents inter-
act with the cytosol through nuclear pores. The single membrane of the endoplasmic
reticulum is highly convoluted. Its extensive network in eukaryotic cells is involved
in the synthesis of transmembrane and secreted proteins and of lipids for many
membranes.

In this section, we explore the structure of biological membranes. In the remaining
sections of this chapter, we discuss the properties and functions of biological membranes.

Citral

CH 2 OH

Bactoprenol
(Undecaprenyl alcohol)

Juvenile hormone I

Limonene

(e)

▲ Figure 9.19

Some isoprenoids. Note the isoprene unit
(red) in bactoprenol.

A. Lipid Bilayers

We saw earlier that detergents in aqueous solutions can spontaneously form monolay-
ers or micelles (Section 2.4). Like detergents, amphipathic glycerophospholipids and
glycosphingolipids can form monolayers under some conditions. In cells, these lipids do
not pack well into micelles but rather tend to form lipid bilayers (Figure 9.20). Lipid bi-
layers are the main structural component of all biological membranes, including
plasma membranes and the internal membranes of eukaryotic cells. The noncovalent

270 CHAPTER 9 Lipids and Membranes

BOX 9.3 GREGOR MENDEL AND GIBBERELLINS

Gregor Mendel studied seven traits in order to come up with
the basic laws of heredity. One of the traits was stem length
(. Le/le ). The Le gene has been cloned and sequenced (Lester et
al., 1997). It encodes the enzyme gibberellin 3/3- hydroxylase,
an enzyme required for the synthesis of the terpenoid gib-
berellin GA1. The production of gibbberellin GA1 by the
normal gene stimulates growth producing a tall pea plant.

The mutant gene produces a less active enzyme that synthe-
sizes less hormone and plants homozygous for the mutant al-
lele (le) are short.

The mutation is a single nucleotide substitution that
converts an alanine codon into a threonine codon (A229T).

Another one of Mendel’s seven traits is described in Box. 15.3.

► The stem length mutation. Tall plants (left) are
normal. Mutations in the stem length gene ( Le )
produce short plants (right).

▲ Figure 9.20

Membrane lipid and bilayer, (a) An amphipathic
membrane lipid, (b) Cross-section of a lipid
bilayer. The hydrophilic head groups (blue)
of each leaflet face the aqueous medium,
and the hydrophobic tails (yellow) pack
together in the interior of the bilayer.

interactions among lipid molecules in bilayers make membranes flexible and allow
them to self-seal. Triacylglycerols, which are very hydrophobic rather than amphipathic,
cannot form bilayers and cholesterol, although slightly amphipathic, does not form
bilayers by itself.

A lipid bilayer is typically about 5 to 6 nm thick and consists of two sheets, or
monolayers (also called leaflets). In each sheet, the polar head groups of amphipathic
lipids are in contact with the aqueous medium and the nonpolar hydrocarbon tails
point toward the interior of the bilayer (Figure 9.20).

The spontaneous formation of lipid bilayers is driven by the hydrophobic interac-
tions (Section 2.5D). When lipid molecules associate, the entropy of the solvent mole-
cules increases and this favors formation of the lipid bilayer.

B. Three Classes of Membrane Proteins

Cellular and intracellular membranes contain specialized membrane-bound proteins.
These proteins are divided into three classes based on their mode of association with the
lipid bilayer: integral membrane proteins, peripheral membrane proteins, and lipid
anchored membrane proteins (Figure 9.21).

Integral membrane proteins , also referred to as transmembrane proteins, contain
hydrophobic regions embedded in the hydrophobic core of the lipid bilayer. Integral
membrane proteins usually span the bilayer completely, with one part of the protein ex-
posed on the outer surface and one part exposed on the inner surface. Some integral
membrane proteins are anchored by only a single membrane-spanning portion of the
polypeptide chain, whereas other membrane proteins have several transmembrane seg-
ments connected by loops at the membrane surface. The membrane-spanning segment
is often an a helix containing approximately 20 amino acid residues.

One of the best characterized integral membrane proteins is bacteriorhodopsin
(Figure 9.22a). This protein is found in the cytoplasmic membrane of the halophilic
(salt-loving) bacterium Halobacterium halobium , where it helps harness light energy
used in the synthesis of ATP. Bacteriorhodopsin consists of a bundle of seven a helices.
The exterior surface of the helical bundle is hydrophobic and interacts directly with
lipid molecules in the membrane. The interior surface contains charged amino acid side
chains that bind the pigment molecule. Bacteriorhodopsin is one of several a-helical
membrane proteins whose structures are known in detail. These a-helix bundle

9.8 Biological Membranes 271

protein protein

▲ Figure 9.21

Structure of a typical eukaryotic plasma

membrane. A lipid bilayer forms the basic matrix of biological membranes, and proteins (some of which are glycoproteins) are associated with it in
various ways. The oligosaccharides of glycoproteins and glycolipids face the extracellular space.

proteins make up one of the two major classes of integral membrane proteins. The
other class is the /3-barrel proteins (see below).

In the absence of data on three-dimensional structure, the presence of transmem-
brane a-helical regions in membrane proteins can often be predicted by searching
amino acid sequences for regions that are hydrophobic (i.e., that have high hydropathy
values) (Section 3.2G) and a tendency to be present in a-helices (Section 4.4). Various
prediction algorithms have been developed over the years and they are currently able to
detect 70% of known transmembrane a-helices. These predictions are important be-
cause it is still very difficult to crystallize membrane proteins in order to determine their
true structure.

v Figure 9.22

Integral membrane proteins, (a) Bacteri-
orhodopsin: seven membrane-spanning a
helices, connected by loops, form a bundle
that spans the bilayer. The light-harvesting
prosthetic group is shown in yellow. [PDB
1FBB]. (b) Porin FhuA from Escherichia
coli\ this porin forms a channel for the pas-
sage of protein-bound iron into the bac-
terium. The channel is formed from 22
antiparallel ft strands that form a /3-barrel.
[PDB 1BY3].

272 CHAPTER 9 Lipids and Membranes

Protein folding is another example of
an entropically driven assembly reac-
tion (Section 4.1 1 A).

We consider the functions of some of
these membrane proteins later in this
chapter. We will also encounter mem-
brane proteins in other chapters,
including those on membrane-associated
electron transport (Chapter 14), photo-
synthesis (Chapter 15), and protein
synthesis (Chapter 22).

The function of bacteriorhodopsin is
described in Section 15.2.

Some prenyl-decorated proteins will be
encountered in the discussion of signal
transduction (Section 9.12).

Many integral membrane proteins have a (3 barrel fold (Figure 4.23b). The exterior
surface of the /3 strands contacts the membrane lipids and the center of the barrel often
serves as a pore or channel for passing molecules from one side of the membrane to the
other. The E. coli porin, FhuA, is a typical example of this type of integral membrane
protein (Figure 9.22b).

Peripheral membrane proteins are associated with one face of the membrane
through charge-charge interactions and hydrogen bonding with integral membrane
proteins or with the polar head groups of membrane lipids. Peripheral membrane pro-
teins are more readily dissociated from membranes by changes in pH or ionic strength.

Lipid anchored membrane proteins are tethered to a membrane through a covalent
bond to a lipid anchor. In the simplest lipid anchored membrane proteins, an amino
acid side chain is linked by an amide or ester bond to a fatty acyl group, often from
myristate or palmitate. The fatty acid is inserted into the cytoplasmic leaflet of the bi-
layer, anchoring the protein to the membrane (Figure 9.23a). Proteins of this type are
found in viruses and eukaryotic cells.

Other lipid anchored membrane proteins are covalently linked to an isoprenoid
chain (either 15- or 20-carbon) through the sulfur atom of a cysteine residue at or near
the C-terminus of the protein (Figure 9.23b). These prenylated proteins are found on the
cytoplasmic face of both plasma membranes and intracellular membranes.

Many eukaryotic lipid anchored proteins are linked to a molecule of glycosylphos-
phatidylinositol (Figure 9.23c). The membrane anchor is the 1,2-diacylglycerol portion
of the glycosylphosphatidylinositol. A glycan of varied composition is attached to the
inositol by a glucosamine residue, a mannose residue links the glycan to a phospho-
ethanolamine residue, and the C-terminal a-carboxyl group of the protein is linked to the
ethanolamine by an amide bond. Over 100 different proteins are known to be associated
with membranes by a glycosylphosphatidylinositol anchor. These proteins have a variety
of functions and they are present only in the outer monolayer of the plasma membrane.
They are found in the cholesterol-sphingolipid raffs described in Section 9.9.

All three types of lipid anchors are covalently linked to amino acid residues post-
translationally, that is, after the protein has been synthesized. Like integral membrane
proteins, most lipid anchored proteins are permanently associated with the membrane,
although the proteins themselves do not interact with the membrane. Once released by
treatment with phospholipases, the proteins behave like soluble proteins.

BOX 9.4 NEW LIPID VESICLES, OR LIPOSOMES

Synthetic vesicles (often called liposomes) consisting of phos-
pholipid bilayers that enclose an aqueous compartment can
be formed in the laboratory. In order to minimize unfavorable
contact between the hydrophobic edge of the bilayer and the
aqueous solution, lipid bilayers tend to close up to form these
spherical structures. The vesicles are generally quite stable and
impermeable to many substances. Liposomes whose aqueous
inner compartment contains drug molecules can be used to
deliver drugs to particular tissues in the body, provided that
specific targeting proteins are present in the liposome mem-
brane. Synthetic bilayers are an important experimental tool
in the investigation of cellular membranes. An example of
such an experiment is described in Box. 15.3.

Lipid

bilayer

Aqueous

solution

1 V

► Schematic cross-section of a lipid vesicle, or liposome. The bilayer is
made up of two leaflets. In each leaflet, the polar head groups of the
amphipathic lipids extend into the aqueous medium and the nonpolar
hydrocarbon tails point inward and are in van der Waals contact with
each other.

Enclosed

aqueous

compartment

9.8 Biological Membranes 273

Phospho-

ethanolamine

residue

Outer

leaflet

Inner

leaflet

◄ Figure 9.23

Lipid anchored membrane proteins attached to the plasma membrane. The three
types of anchors can be found in the same membrane, but they do not form a
complex as shown here, (a) A fatty acyl anchored protein, (b) A prenyl
anchored membrane protein. Note that fatty acyl and prenyl anchored mem-
brane proteins can also occur on the cytoplasmic (outer) leaflet of intracellular
membranes, (c) Protein anchored by glycosy I phosphatidyl i nositol . Shown here
is the variant surface glycoprotein of the parasitic protozoan Trypanosoma
brucei. The protein is covalently bound to a phosphoethanolamine residue,
which in turn is bound to a glycan. The glycan (blue) includes a mannose
residue to which the phosphoethanolamine residue is attached and a glu-
cosamine residue that is attached to the phosphoinositol group (red) of
phosphatidylinositol. Abbreviations: GlcN, glucosamine; Ins, inositol; Man,
mannose.

The total number of membrane proteins in a typical cell isn’t known for certain
but they are likely to represent a significant fraction of the proteome. In E. coli , for ex-
ample, there appear to be roughly 1000 membrane proteins of all types. Since the total
number of proteins is about 4000 (Chapter 4), membrane proteins account for about
25% of the total. This fraction is probably higher in multicellular eukaryotes because
there are many more membrane proteins involved in cell-cell interactions and intracel-
lular signaling.

Different membranes have different proteins (and lipids). In some cases a cell or
compartment is enclosed by a double membrane consisting of two separate lipid bilayers
(Figure 9.24). In the case of mitochondria and E. coli , the inner membranes have many
more membrane proteins than the outer membranes.

Figure 9.24 ►

Double membrane of mitochondria and many bacteria. The plasma membrane of most eukaryotic cells
is a single lipid bilayer. Within eukaryotic cells the nucleus and major organelles such as mitochon-
dria (top right) are bounded by double membranes. In bacteria, the gram-negative bacteria have a
double membrane consisting of an inner and outer lipid bilayer as shown for E. coli (bottom right).
It’s not surprising that mitochondria (and chloroplasts) have a double membrane since they are de-
rived from gram-negative bacteria that use the double membrane as part of the energy-producing
mechanism of electron transport and ATP synthesis (Chapter 14).

274 CHAPTER 9 Lipids and Membranes

BOX 9.5 SOME SPECIES HAVE UNUSUAL LIPIDS IN THEIR MEMBRANES

Many species have unusual lipids in some of their mem-
branes. The unusual lipids are sometimes confined to genera
or families and sometimes entire orders share some distinc-
tive lipid compositions. Within the eukaryotes, there are
some lipids found only in some classes of animals and not
others or in some classes of plants and not others. There are
even distinctive lipid compositions in some entire kingdoms
such as plants, animals, or fungi.

Prokaryotes are a very diverse group with many varieties
of lipids. Major groups such as cyanobacteria, mycoplasma,
and gram positive bacteria, can have quite characteristic lipid
compositions in their membranes.

The archaebacteria (or Archaea) have glycerophospholipids
that are quite unusual and distinctive. The glycerol phosphate

backbone in archaebacterial glycerophospholipids is sn-glycerol-
1 -phosphate, a stereoisomer of the one found in other species
(sn-glycerol-3-phosphate). (see Box 16.1) The hydrocarbon
chains are attached to the glycerol backbone via ether linkages,
not ester linkages, and the hydrocarbon chains in archaebacteria
are often isoprenoid derivatives, not fatty acid derivatives.

There are a few species of gram-negative bacteria that
have mixtures of ether and ester linkages in their lipids but
unusual lipid composition of archaebacteria argues strongly
in favor of classifying them as a distinctive monophyletic
group. As mentioned earlier (Section 1.5), some scientists
argue that the distinctiveness of archaebacteria justifies creat-
ing a third domain of life but the current view favors a more
complex web of life perspective.

Ether linkage

sn-G-3-P backbone

Ester linkage Fatty acid chain

◄ Comparison of typical bacterial
and archaebacterial glycero phos-
pholipids.

Archaea

Bacteria

C. The Fluid Mosaic Model of Biological Membranes

A typical biological membrane contains about 25% to 50% lipid and 50% to 75% protein
by mass. Carbohydrates are present as components of glycolipids and glycoproteins.
The lipids are a complex mixture of phospholipids, glycosphingolipids (in ani-
mals), and cholesterol (in some eukaryotes). Cholesterol and some other lipids that do
not form bilayers by themselves (about 30% of the total) are stabilized in a bilayer
arrangement by the other 70% of lipids in the membrane (see next section).

The compositions of biological membranes vary considerably among species and
even among different cell types in multicellular organisms. For example, the myelin
membrane that insulates nerve fibers contains relatively little protein. In contrast, the
inner mitochondrial membrane is rich in proteins reflecting its high level of meta-
bolic activity. The plasma membrane of red blood cells is also exceptionally rich in
proteins.

Each biological membrane has a characteristic lipid composition, in addition to
having a characteristic lipid to protein ratio. Membranes in brain tissue, for example,
have a relatively high content of phosphatidylserines whereas membranes in heart and
lung cells have high levels of phosphatidylglycerols and sphingomyelins, respectively.
Phosphatidylethanolamines constitute nearly 70% of the inner membrane lipids of
E. coli cells. The outer membranes of gram-negative bacteria contain lipopolysaccharides.

9.9 Membranes Are Dynamic Structures

275

In addition to being distributed differentially among different tissues, phospho-
lipids are also distributed asymmetrically between the inner and outer monolayers of a
single biological membrane. In mammalian cells, for example, 90% of the sphingomyelin
molecules are in the outer surface of the plasma membrane. Phosphatidylserines are
also asymmetrically distributed in many cells, with 90% of the molecules in the cyto-
plasmic monolayer.

A biological membrane is thicker than a lipid bilayer — typically 6 to 10 nm thick.
The fluid mosaic model proposed in 1972 by S. Jonathan Singer and Garth L. Nicolson is
still generally valid for describing the arrangement of lipid and protein within a mem-
brane. According to the fluid mosaic model, the membrane is a dynamic structure in
which both proteins and lipids can rapidly and randomly diffuse laterally or rotate
within the bilayer. Membrane proteins are visualized as icebergs floating in a highly
fluid lipid bilayer sea (Figure 9.21). (Actually, some proteins are immobile and some
lipids have restricted movement.)

KEY CONCEPT

Membranes consist of a lipid bilayer and
embedded proteins. Lipids and proteins
can diffuse rapidly within the membrane.

9.9 Membranes Are Dynamic Structures

The lipids in a bilayer are in constant motion giving lipid bilayers many of the proper-
ties of fluids. A lipid bilayer can therefore be regarded as a two-dimensional solution.
Lipids undergo several types of molecular motion within bilayers. The rapid movement
of lipids within the plane of one monolayer is an example of two-dimensional lateral
diffusion. A phospholipid molecule can diffuse from one end of a bacterial cell to the
other (a distance of about 2 \x m) in about 1 second at 37°C.

In contrast, transverse diffusion (or flip-flop) is the passage of lipids from one
monolayer of the bilayer to the other. Transverse diffusion is much slower than lateral
diffusion (Figure 9.25). The polar head of a phospholipid molecule is highly solvated
and must shed its solvation sphere and penetrate the hydrocarbon interior of the bilayer
in order to move from one leaflet to the other. The energy barrier associated with this
movement is so high that transverse diffusion of phospholipids in a bilayer occurs at
about one-billionth the rate of lateral diffusion. The very slow rate of transverse diffu-
sion of membrane lipids is what allows the inner and outer layers of biological mem-
branes to maintain different lipid compositions.

All cells synthesize new membrane by adding lipids and protein to preexisting
membranes. As the plasma membrane is extended, the cell increases in size. Eventually
the cell will divide and each daughter cell will inherit a portion (usually half) of the
parental membranes. Internal membranes are extended and divide in the same manner.

In bacteria, lipid molecules are usually added to the cytoplasmic side of the lipid bi-
layer. Lipid asymmetry is generated by preferentially adding newly synthesized lipids to

You might have inherited lipid mole-
cules from your grandmother! (see
Problem 18).

(a) Lateral diffusion

(b) Transverse diffusion

◄ Figure 9.25

Diffusion of lipids within a bilayer, (a) Lateral
diffusion of lipids is relatively rapid.

(b) Transverse diffusion, or flip-flop, of lipids
is very slow.

276 CHAPTER 9 Lipids and Membranes

Human cell Mouse cell

Red fluorescent Green fluorescent
markers markers

Immediately after fusion,
fluorescent markers remain localized.

Within 40 minutes, fluorescent
markers appear to be randomly
distributed over the entire surface.

▲ Figure 9.26

Diffusion of membrane proteins. Human cells
whose membrane proteins had been labeled
with a red fluorescent marker were fused
with mouse cells whose membrane proteins
had been labeled with a green fluorescent
marker. The initially localized markers be-
came dispersed over the entire surface of
the fused cell within 40 minutes.

only one of the monolayers. Since transverse diffusion is so slow, these newly synthesized
molecules will not spread to the outer layer of the plasma membrane. This accounts for
the enrichment of some types of lipids in the inner layer. Lipid asymmetry can also be
generated and maintained by the activity of membrane-bound flipases and flopases — en-
zymes that use the energy of ATP to move specific phospholipids from one monolayer to
the other. The activity of these enzymes accounts for the enrichment of certain types of
phospholipid in the outer layer. Eukaryotic cells make their membrane lipids in an asym-
metric arrangement in the endoplasmic reticulum or the Golgi apparatus. The membrane
fragments flow from these organelles — retaining the asymmetry — to other membranes.

In 1970, L. D. Frye and Michael A. Edidin devised an elegant experiment to
test whether membrane proteins diffuse within the lipid bilayer. Frye and Edidin
fused mouse cells with human cells to form heterokaryons (hybrid cells). By using red
fluorescence-labeled antibodies that specifically bind to certain proteins in human
plasma membranes and green fluorescence-labeled antibodies that specifically bind to
certain proteins in mouse plasma membranes, they observed the changes in the distri-
bution of membrane proteins over time by immunofluorescence microscopy. The
labeled proteins were intermixed within 40 minutes after cell fusion (Figure 9.26).
This experiment demonstrated that at least some membrane proteins diffuse freely
within biological membranes.

A few membrane proteins move laterally very rapidly but the majority of mem-
brane proteins diffuse about 100 to 500 times more slowly than membrane lipids.
The diffusion of some proteins is severely restricted by aggregation or by attach-
ment to the cytoskeleton just beneath the membrane surface. Relatively immobile
membrane proteins may act as fences or cages, restricting the movement of other
proteins. The limited diffusion of membrane proteins produces protein patches,
or domains — areas of membrane whose composition differs from that of the
surrounding membrane.

The distribution of membrane proteins can be visualized by freeze- fracture elec-
tron microscopy. In this technique, a membrane sample is rapidly frozen to the temper-
ature of liquid nitrogen and then fractured with a knife. The membrane splits between
the leaflets of the lipid bilayer where the intermolecular interactions are weakest
(Figure 9.27a). Ice is evaporated in a vacuum and the exposed internal surface of the
membrane is then coated with a thin film of platinum to make a metal replica for ex-
amination in an electron microscope. Membranes that are rich in membrane proteins
contain pits and bumps indicating the presence of proteins. In contrast, membranes
that contain no proteins are smooth. Figure 9.27b shows the bumpy surface of the
inner monolayer of a red blood cell membrane exposed by removal of the outer layer.

The fluid properties of lipid bilayers depend on the flexibility of their fatty acyl
chains. Saturated acyl chains are fully extended at low temperatures forming a crystalline
array with maximal van der Waals contact between the chains. When the lipid bilayer is
heated, a phase transition analogous to the melting of a crystalline solid occurs. The acyl
chains of lipids in the resulting liquid crystalline phase are relatively disordered and
loosely packed. During the phase transition, the thickness of the bilayer decreases
by about 15% as the hydrocarbon tails become less extended because of rotation around
C — C bonds (Figure 9.28). Bilayers composed of a single type of lipid undergo phase
transition at a distinct temperature called the phase-transition temperature. When the
lipids contain unsaturated acyl chains, the hydrophobic core of the bilayer is fluid well
below room temperature (23°C). Biological membranes, which contain a heterogeneous
mixture of lipids, change gradually from the gel to the liquid crystalline phase, typically
over a temperature range of 10° to 40°C. Phase transitions in biological membranes can
be localized so fluid- and gel-phase regions can coexist at certain temperatures.

The structure of a phospholipid has dramatic effects on its fluidity and phase-transition
temperature. As we saw in Section 9.2, the hydrocarbon chain of a fatty acid with a cis
double bond has a kink that disrupts packing and increases fluidity. Incorporating an un-
saturated fatty acyl group into a phospholipid lowers the phase-transition temperature.
Changes in membrane fluidity affect the membrane transport and catalytic functions of
membrane proteins so many organisms maintain membrane fluidity under different con-
ditions by adjusting the ratio of unsaturated to saturated fatty acyl groups in membrane

9.10 Membrane Transport 277

(a)

Inner

leaflet

Outer

leaflet

(b)

Outer Inner
surface leaflet

▲ Figure 9.27

Freeze fracturing a biological membrane.

(a) Splitting the lipid bilayer along the interface of the two leaflets. A platinum replica of the exposed internal surface is examined in an electron mi-
croscope. Membrane proteins appear as protrusions or cavities in the replica, (b) Electron micrograph of a freeze-fractured erythrocyte membrane.
The bumps on the inner membrane surface show the locations of membrane proteins.

lipids. For example, when bacteria are grown at low temperatures, the proportion of un-
saturated fatty acyl groups in membranes increases. Goldfish adapt to the temperature of
the water in which they swim: as the environmental temperature drops, there is a rise in
unsaturated fatty acids in goldfish intestinal membranes and whole brain. The lower
melting point and greater fluidity of unsaturated fatty acyl groups preserve membrane
fluidity allowing membrane processes to continue at colder temperatures.

Cholesterol accounts for 20% to 25% of the mass of lipids in a typical mammalian
plasma membrane and significantly affects membrane fluidity. When the rigid choles-
terol molecules intercalate between the hydrocarbon chains of the membrane lipids, the
mobility of fatty acyl chains in the membrane is restricted and fluidity decreases at high
temperatures (Figure 9.29). Cholesterol disrupts the ordered packing of the extended
fatty acyl chains and thereby increases fluidity at low temperatures. Cholesterol in ani-
mal cell membranes thus helps maintain fairly constant fluidity despite fluctuations in
temperature or degree of fatty acid saturation.

Cholesterol tends to associate with sphingolipids because they have long saturated
fatty acid chains. The unsaturated chains of most glycerophospholipids produce kinks
that don’t easily accommodate cholesterol molecules in the membrane. Because of this
preferential association, mammalian membranes consist of patches of cholesterol/
sphingolipids regions surrounded by regions that have very little cholesterol. These
patches are called lipid rafts. Certain membrane proteins may preferentially associate
with lipid rafts. Thus, some membrane proteins may also have a patch-like distribution
on the cell surface. Membrane proteins are thought to play an important role in main-
taining the integrity of lipid rafts.

Ordered gel Disordered liquid

phase crystalline phase

▲ Figure 9.28

Phase transition of a lipid bilayer. In the or-
dered gel state, the hydrocarbon chains are
extended. Above the phase-transition temper-
ature, rotation around C — C bonds disorders
the chains in the liquid crystalline phase.

9.10 Membrane Transport

Plasma membranes physically separate a living cell from its environment. In addition,
within both prokaryotic and eukaryotic cells there are membrane-bound compartments.
The nucleus and mitochondria are obvious examples in eukaryotes.

278 CHAPTER 9 Lipids and Membranes

(a)

▲ Goldfish adapt to water temperature, (a) These
goldfish (carp, Carassius auratus ) have
adapted to the water temperature in Kyoto,
Japan, by adjusting the lipid composition of
their membranes, (b) These Goldfish® do
not adapt well to any water temperature.

▲ Figure 9.29

Model of a lipid membrane. Cholesterol mole-
cules (green) are packed between phospholipid
fatty acid chains (grey).

Membranes are selectively permeable barriers that restrict the free passage of
most molecules. As a general rule, the permeability of molecules is related to their hy-
drophobicity and their tendency to dissolve in organic solvents. Thus, hexanoic acid,
acetic acid, and ethanol are able to move across membranes quite readily. They have
high permeability coefficients (Figure 9.30). Water, despite its strong polar character,
is able to diffuse freely across lipid bilayers although, as the permeability coefficient
indicates, its movement is still greatly restricted compared to organic solvents like
hexanoic acid.

Small ions like Na + , K + , and CP have very low permeability coefficients. They are
unable to diffuse across a membrane because the hydrophobic core of the lipid bilayer
presents an almost impenetrable barrier to most polar or charged species. H + ions have
a much higher permeability coefficient although membranes still act as an effective
barrier to protons.

As mentioned above, very hydrophobic molecules and some small uncharged mol-
ecules can move through biological membranes. Water, oxygen, and other small mole-
cules must also be able to enter all cells and move freely between compartments inside
eukaryotic cells even if they are not able to diffuse as quickly across membranes. Larger
molecules, such as proteins and nucleic acids, have to be transported across mem-
branes, including the membranes between compartments. Living cells move molecules
across membranes using transport proteins (sometimes called pores, carriers, perme-
ases, or pumps) and they transport macromolecules by endocytosis or exocytosis.

Nonpolar gases, such as 0 2 and C0 2 , and hydrophobic molecules, such as steroid
hormones, lipid vitamins, and some drugs, enter and leave the cell by diffusing through
the membrane moving from the side with the higher concentration to the side with the
lower concentration. The rate of movement depends on the difference in concentra-
tions, or the concentration gradient, between the two sides. Diffusion down a concen-
tration gradient (i.e., downhill diffusion) is a spontaneous process driven by an increase
in entropy and therefore a decrease in free energy (see below).

The traffic of other molecules and ions across membranes is mediated by three
types of integral membrane proteins: channels and pores, passive transporters, and ac-
tive transporters. These transport systems differ in their kinetic properties and energy
requirements. For example, the rate of solute movement through pores and channels
may increase with increasing solute concentration but the rate of movement through
passive and active transporters may approach a maximum as the solute concentration
increases (i.e., the transport protein becomes saturated). Some types of transport re-
quire a source of energy (Section C). The characteristics of membrane transport are
summarized in Table 9.3. In this section, we describe the different membrane transport
systems, as well as endocytosis and exocytosis.

A. Thermodynamics of Membrane Transport

Recall from Chapter 1 (Section 1.4C) that the actual Gibbs free energy change of a reac-
tion is related to the standard Gibbs free energy change by the equation

[C][D]

A ^reaction — reaction + RT In jgj (9.1)

where AG°' react i on represents the standard Gibbs free energy change for the reaction,
[C] [D] represents the concentrations of the products, and [A] [B] represents the con-
centration of the reactants. The Gibbs free energy change associated with membrane
transport depends only on the concentrations of the molecules on either side of the
membrane.

For any molecule, A, the concentration on the inside of the membrane is [ Aj n ] and
the concentration outside is [A out ] . The Gibbs free energy change associated with trans-
porting molecules of A is

[A in ] [A in ]

AG transport = RT In-^A = 2.303 RT (9.2)

l/VDUtJ l/VDUtJ

9.10 Membrane Transport 279

Table 9.3 Characteristics of different types of membrane transport

Protein

carrier

Saturable

with

substrate

Movement
relative to
concentration
gradient

Energy input
required

Simple diffusion

Down

Channels and pores

Yes

Down

Passive transport

Yes

Down

Active transport

Primary

Yes

Yes (direct source)

Secondary

Yes

Yes (ion gradient)

If the concentration of A inside the cell is much less than the concentration of A out-
side the cell then AG transport will be negative and the flow of A into the cell will be ther-
modynamically favored. For exmple, if [A an ] = 1 mM and [A out ] = 100 mM, then at 25°C

TA- 1

AGtransport = 2.303 RT \ogj-^ = 2.303 X 8.325 X 298 X (-2)

L oud (9 . 3)

= -1 1 .4 kj mol -1

Under these conditions, molecules of solute A will tend to flow into the cell in order to
reduce the concentration gradient. Flow in the opposite direction is thermodynamically
unfavorable since it is associated with a positive Gibbs free energy change (AG transport =
+ 1 1.4 kj mol -1 for molecules moving from the inside of the cell to the outside).

Equation 9.2 only applies to uncharged molecules. In the case of ions, the Gibbs free
energy change has to include a factor that takes into account the charge difference across
a biological membrane. Most cells selectively export cations so the inside of a cell is neg-
atively charged with respect to the outside. The charge difference across the membrane is

AT' = T' in - T'out (9.4)

where AT' is called the membrane potential (in volts). The Gibbs free energy change
due to this electric potential is

AG = zFAT' (9.5)

where z is the charge on the molecule being transported (e.g., +1,-1, +2, —2, etc.) and F
is Faradays’s constant (96,485 JV -1 mol -1 ). Since the inside of the cell is negatively
charged, the import of cations such as Na© and K© is thermodynamically favored by
the membrane potential. The export of cations must be coupled to an energy-producing
reaction since it is associated with a positive Gibbs free energy change.

Both the chemical (concentration) and electric (charge) effects have to be consid-
ered, for any transport process involving charged molecules. Thus,

AG transport = 2.303 RT log + zFW (9 . 6)

B. Pores and Channels

Pores and channels are transmembrane proteins with a central passage for ions and
small molecules. (Usually, the term pore is used for bacteria and channel for animals.)
Solutes of the appropriate size, charge, and molecular structure can move rapidly

Permeability
coefficient
(cm s 1 )

= ^ Hexanoic acid

10- 1 -

10- 2

10 3

Acetic acid

Water

Ethanol

10“ 4

10 5 -

10“ 6 -

Indole

H +

Glycerol, Urea

10- 7

10- 8

Tryptophan

Glucose

10- 9 -=

10- 11 -E

10- 12 -= — Na +

1 0 -!3 J

▲ Figure 9.30

Permeability coefficients of various molecules.

Molecules with high permeability coeffi-
cients (top) are able to diffuse unaided
across a membrane.

KEY CONCEPT

For a given solute, the Gibbs free energy
change of transport depends on both the
membrane potential and solute concen-
trations on either side of the membrane.

The importance of Equation 9.6 will
become apparent when we describe
chemiosmotic theory (Section 14.3).

280 CHAPTER 9 Lipids and Membranes

a Membrane potential. In most cases the in-
side of a cell or membrane compartment is
negative with respect to the outside and the
membrane potential (A\| /) is negative.

O o° Q o

°o

▲ Figure 9.31

Membrane transport through a pore or channel.

A central passage allows molecules and ions
of the appropriate size, charge, and geometry
to traverse the membrane in either direction.

through the passage in either direction by diffusing down a concentration gradient
(Figure 9.31). This process requires no energy. In general, the rate of movement of
solute through a pore or channel is not saturable at high concentrations. For some
channels, the rate may approach the diffusion controlled limit.

The outer membranes of some bacteria are rich in porins, a family of pore proteins
that allow ions and many small molecules to gain access to specific transporters in the
plasma membrane. Similar channels are found in the outer membranes of mitochon-
dria. Porins are usually only weakly solute-selective. They can act as sieves that are per-
manently open or they can be regulated by the concentration of solutes. In contrast,
plasma membranes also contain many channel proteins that are highly specific for cer-
tain ions and they open or close in response to a specific signal.

Aquaporin is an integral membrane protein that acts as a pore for water molecules.
The channel through the middle of the protein will allow for passage of water molecules
and other small uncharged molecules but it blocks passage of any charged molecules or
large molecules. This channel is larger on the outside surface but narrows to a much
smaller channel on the cytoplasmic side as shown for yeast aquaporin in Figure 9.32.
Aquaporins are common in all species. They are required in cells where the rapid uptake
of water is necessary because the rate of diffusion of water across the membrane is too
slow. This is an example of a simple, somewhat specific, porin. It was discovered by
Peter Agre, who received the Nobel Prize in Chemistry in 2003.

Cor A is the primary Mg 2+ pump in prokaryotic cells. It is highly selective for Mg 2+
and permits the import of Mg 2+ against a concentration gradient in response to the
membrane potential. Positively charged ions “want” to flow into cells and the CorA pore
allows passage of Mg 2+ but not other ions. Mg 2+ is essential for many cell functions. The
rate of influx is regulated by the large cytoplasmic domain of CorA (Figure 9.33). It
binds Mg 2+ ions and when a sufficient number have bound, the pore is closed. Thus, in-
flux of Mg 2+ is controlled by the cytoplasmic concentration.

Membranes of nerve tissues have gated (i.e., controlled) potassium channels that
selectively allow rapid outward transport of potassium ions. These channels permit
K© ions to pass through the membrane at least 10,000 times faster than the smaller
Na© ions. Crystallographic studies have shown that the potassium channel has a wide
mouth (like a funnel) containing negatively charged amino acids to attract cations and
repel anions. Hydrated cations are directed electrostatically to an electrically neutral
constriction of the pore called the selectivity filter. Potassium ions rapidly lose some of
their water of hydration and pass through the selectivity filter. Sodium ions apparently
retain more water of hydration and therefore transit the filter much more slowly. The
remainder of the channel has a hydrophobic lining. Based on comparisons of amino
acid sequences, the general structural properties of the potassium channel seem to also
apply to other types of channels and pores. Roderick MacKinnon shared the 2003
Nobel Prize in Chemistry with Peter Agre. MacKinnnons work focused mainly on
potassium channels.

C. Passive Transport and Facilitated Diffusion

Pore and channel proteins are examples of passive transport where the Gibbs free energy
change for transport is negative and transport from one side of the membrane to the
other is a spontaneous process. In active transport (see below), the solute moves against
a concentration gradient and/or a charge difference. Active transport must be coupled
to an energy-producing reaction in order to overcome the unfavorable Gibbs free en-
ergy change for unassisted transport. The simplest membrane transporters — whether
active or passive — carry out uni port; that is, they carry only a single type of solute across
the membrane (Figure 9.34a). Many transporters carry out the simultaneous transport
of two different solute molecules. The process is called symport if both solutes are

◄ Figure 9.32

Fungal aquaporin. Aquaporin is an integral membrane protein with an a-helix bundle domain. The
water channel (green dots) is open on the exterior surface and narrows to a tiny passage on the
cytoplasmic side. [Pichia pastoris PDB 2W2E]

9.10 Membrane Transport

281

Figure 9.33 ▲

CorA, a magnesium pump. CorA is the prokaryotic magnesium pump. Mg 2+ ions bind on the exterior
surface and are transported through a highly selective channel in response to the membrane po-
tential. The cytoplasmic domain binds Mg 2+ ions and closes the pore in response to high internal
concentrations of Mg 2+ . This is the Thermotoga maritima version with each of the fire subunits in
a different color. [PDB 2HN2]

transported in the same direction, (Figure 9.34b). If they are transported in opposite di-
rections, the process is anti port (Figure 9.34c).

Passive transport includes simple diffusion across a membrane. When pores, chan-
nels, and transporters are involved, we call the process facilitated diffusion. Facilitated
diffusion is still an example of passive transport since it does not require an energy source.
The transport protein accelerates the movement of solute down its concentration gradi-
ent, or charge gradient, a process that would occur very slowly by diffusion alone. In this
case, transport proteins are similar to enzymes because they increase the rate of a process
that is thermodynamically favorable. For a simple passive uniport system, the initial rate
of inward transport, like the initial rate of an enzyme- catalyzed reaction, depends on the
external concentration of substrate. The equation describing this dependence is analogous
to the Michaelis-Menten equation for enzyme catalysis (Equation 5.14).

Knax[S]out

= K , rcn ( 9 - 7 >

Ktr + LMout

where v 0 is the initial rate of inward transport of the substrate at an external concentra-
tion [S] out , V mSK is the maximum rate of transport of the substrate, and K tr is a constant
analogous to the Michaelis constant (K m ) (i.e., K tr is the substrate concentration at
which the transporter is half-saturated). The lower the value of K tr , the higher the affin-
ity of the transporter for the substrate. The rate of transport is saturable, approaching a
maximum value at a high substrate concentration (Figure 9.35).

As substrate accumulates inside the cell, the rate of outward transport increases
until it equals the rate of inward transport, and [S]j n equals [S] out . At this point, there is
no net change in the concentration of substrate on either side of the membrane, al-
though substrate continues to move across the membrane in both directions.

Models of transport protein operation suggest that some transporters undergo a
conformational change after they bind their substrates. This conformational change al-
lows the substrate to be released on the other side of the membrane; the transporter

v o

▲ Figure 9.34

Types of passive and active transport. Although
the transport proteins are depicted as having
an open central pore, passive and active
transporters actually undergo conformational
changes when transporting their solutes.

(a) Uniport, (b) Symport. (c) Antiport.

282 CHAPTER 9 Lipids and Membranes

▲ Figure 9.35

Kinetics of passive transport. The initial rate of
transport increases with substrate concentration
until a maximum is reached. K tr is the concen-
tration of substrate at which the rate of trans-
port is half-maximal.

then reverts to its original state (Figure 9.36). The conformational change in the trans-
porter is often triggered by binding of the transported species, as in the induced fit of
certain enzymes to their substrates (Section 6.9). In active transport, the conforma-
tional change can be driven by ATP or other sources of energy. Like enzymes, trans-
port proteins can be susceptible to reversible and irreversible inhibition.

D. Active Transport

Active transport resembles passive transport in overall mechanism and kinetic proper-
ties. However, active transport requires energy to move a solute up its concentration
gradient. In some cases, active transport of charged molecules or ions also results in a
charge gradient across the membrane and active transport moves ions against the
membrane potential.

Active transporters use a variety of energy sources, most commonly ATP. Ion-
transporting ATPases are found in all organisms. These active transporters, which in-
clude Na©-K© ATPase, and Ca® ATPase, create and maintain ion concentration gradients
across the plasma membrane and across the membranes of internal organelles.

Primary active transport is powered by a direct source of energy such as ATP or
light. For example, bacteriorhodopsin (Figure 9.22) uses light energy to generate a
transmembrane proton concentration gradient that can be used for ATP formation.
One primary active transport protein, P- glycoprotein, appears to play a major role in
the resistance of tumor cells to multiple chemotherapeutic drugs. Multidrug resist-
ance is a leading cause of failure in the clinical treatment of human cancers. P- Glyco-
protein is an integral membrane glycoprotein (M r 170,000) that is abundant in the
plasma membrane of drug-resistant cells. Using ATP as an energy source, P-glycopro-
tein pumps a large variety of structurally unrelated nonpolar compounds, such as
drugs, out of the cell up a concentration gradient. In this way, the cytosolic drug con-
centration is maintained at a level low enough to avoid cell death. The normal physi-
ological function of P- glycoprotein appears to be removal of toxic hydrophobic com-
pounds in the diet.

Secondary active transport is driven by an ion concentration gradient. The active
uphill transport of one solute is coupled to the downhill transport of a second solute
that was concentrated by primary active transport. For example, in E. coli , electron
flow through a series of membrane-bound oxidation-reduction enzymes generates a
higher extracellular concentration of protons. As protons flow back into the cell down
their concentration gradient, lactose is also transported in, against its concentration
gradient (Figure 9.37). The energy of the proton concentration gradient drives the sec-
ondary active transport of lactose. The symport of H© and lactose is mediated by the
transmembrane protein lactose permease.

In large multicellular animals, secondary active transport is often powered by a
sodium ion gradient. Most cells maintain an intracellular potassium ion concentra-
tion of about 140 mM in the presence of an extracellular concentration of about 5
mM. The cytosolic concentration of sodium ions is maintained at about 5 to 15 mM
in the presence of an extracellular concentration of about 145 mM. These ion con-
centration gradients are maintained by Na©-K© ATPase, an ATP-driven antiport
system that pumps two K© into the cell and ejects three Na© for every molecule of
ATP hydrolyzed (Figure 9.38). Each Na©-K© ATPase can catalyze the hydrolysis of
about 100 molecules of ATP per minute, a significant portion (up to one-third) of the
total energy consumption of a typical animal cell. The Na© gradient that is generated
by Na©-K© ATPase is the major source of energy for secondary active transport of
glucose in intestinal cells. One glucose molecule is imported with each sodium ion
that enters the cell. The energy released by the downhill movement of Na© powers
the uphill transport of glucose.

◄ Figure 9.36

Passive and active transport protein function. The protein binds its specific substrate and then under-
goes a conformational change, allowing the molecule or ion to be released on the other side of the
membrane. Cotransporters have specific binding sites for each transported species.

9.11 Transduction of Extracellular Signals 283

E. Endocytosis and Exocytosis

The transport we have discussed so far occurs by the flow of molecules or ions across an
intact membrane. Cells also need to import and export molecules too large to be trans-
ported via pores, channels, or transport proteins. Prokaryotes possess specialized multi-
component export systems in their plasma and outer membranes that allow them to se-
crete certain proteins (often toxins or enzymes) into the extracellular medium. In
eukaryotic cells, many — but not all — proteins (and certain other large substances) are
moved into and out of the cell by endocytosis and exocytosis, respectively. In both cases,
transport involves formation of a specialized type of lipid vesicle.

Endocytosis is the process by which macromolecules are engulfed by the plasma
membrane and brought into the cell inside a lipid vesicle. Receptor- mediated endo-
cytosis begins with the binding of macromolecules to specific receptor proteins in
the plasma membrane of the cell. The membrane then invaginates, forming a vesicle
that contains the bound molecules. As shown in Figure 9.39, the inside of such a
membrane vesicle is equivalent to the outside of a cell; thus, substances inside the
vesicle have not actually crossed the plasma membrane. Once inside the cell, the
vesicle can fuse with an endosome (another type of vesicle) and then with a lyso-
some. Inside a lysosome, the endocytosed material and the receptor itself can be
degraded. Alternatively, the ligand, the receptor, or both, can be recycled from the
endosome back to the plasma membrane.

Exocytosis is similar to endocytosis except that the direction of transport is re-
versed. During exocytosis, materials destined for secretion from the cell are enclosed
in vesicles by the Golgi apparatus (Section 1.8B). The vesicles then fuse with the
plasma membrane releasing the vesicle contents into the extracellular space. The
zymogens of digestive enzymes are exported from pancreatic cells in this manner
(Section 6.7A).

Lactose H

▲ Figure 9.37

Secondary active transport in Escherichia coli. The

oxidation of reduced substrates (S rec j) generates a
transmembrane proton concentration gradient.

The energy released by protons moving down their
concentration gradient drives the transport of lac-
tose into the cell by lactose permease.

The secretory pathway in eukaryotic
cells is described in Section 22.10.

9.11 Transduction of Extracellular Signals

In order for a cell to interact with its external environment, it must detect molecules
outside of the plasma membrane and convey that information to the inside of the cell.
This process is called signal transduction and it is a very active field of research. In this
section we’ll cover the basic mechanism of the most common signaling pathways. As
you learn more biochemistry, you’ll encounter many variations of these themes.

A. Receptors

The plasma membranes of all cells contain specific receptors that allow the cell to
respond to external chemical stimuli that cannot cross the membrane. For example,

EXTERIOR

[K©] = 5mM

3Na© 2 K©

Na© Glucose

◄ Figure 9.38

Secondary active transport in animals. The

Na©-K© ATPase generates a sodium ion
gradient that drives secondary active trans-
port of glucose in intestinal cells.

284

CHAPTER 9 Lipids and Membranes

▲ Figure 9.39

Electron micrographs of endocytosis. Endocytosis
begins with the binding of macromolecules
to the plasma membrane of the cell. The
membrane then invaginates forming a vesi-
cle that contains the bound molecules. The
inside of the vesicle is topologically equiva-
lent to the outside of the cell.

bacteria can detect certain chemicals in their environment. A signal is passed via a
cell surface receptor to the flagella, causing the bacterium to swim toward a potential
food source. This is called positive chemotaxis. In negative chemotaxis, the bacteria
swim away from toxic chemicals.

In multicellular organisms, stimuli such as hormones , neurotransmitters (sub-
stances that transmit nerve messages at synapses), and growth factors (proteins that
regulate cell proliferation) are produced by specialized cells. These ligands can travel to
other tissues where they bind to and produce specific responses in cells with the appro-
priate receptors on their surfaces. In this section, we see how the binding of water-
soluble ligands to receptors elicits intracellular responses in mammals. These signal
transduction pathways involve adenylyl cyclase, inositol phospholipids, and receptor
tyrosine kinases.

BOX 9.6 THE HOT SPICE OF CHILI PEPPERS

Biochemists now know the mechanism by which spice from
“hot” peppers exerts its action, causing a burning pain. The
active factor in capsaicin peppers is a lipophilic vanilloid
compound called capsaicin.

Capsaicin

A nerve cell protein receptor that responds to cap-
saicin has been identified and characterized. It is an ion
channel and its amino acid sequence suggests that it has
six transmembrane domains. Activation of the receptor
by capsaicin causes the channel to open so that calcium
and sodium ions can flow into the nerve cell and send an

Chili peppers ►

impulse to the brain. The receptor is activated not only by
vanilloid spices but also by rapid increases in temperature. In
fact, the main function of the receptor is detection of heat.

9.11 Transduction of Extracellular Signals 285

External stimulus

PLASMA

MEMBRANE

Second messenger

JJ^DNA binding

Cytoplasmic and nuclear effectors

Cellular response

◄ Figure 9.40

General mechanism of signal transduction
across the plasma membrane of a cell.

A general mechanism for signal transduction is shown in Figure 9.40. A ligand
binds to its specific receptor on the surface of the target cell. This interaction generates a
signal that is passed through a membrane protein transducer to a membrane-bound
effector enzyme. The action of the effector enzyme generates an intracellular second
messenger that is usually a small molecule or ion. The diffusible second messenger car-
ries the signal to its ultimate destination which may be in the nucleus, an intracellular
compartment, or the cytosol. Ligand binding to a cell-surface receptor almost invari-
ably results in the activation of protein kinases. These enzymes catalyze the transfer
of a phosphoryl group from ATP to various protein substrates, many of which help
regulate metabolism, cell growth, and cell division. Some proteins are activated by
phosphorylation, whereas others are inactivated. A vast diversity of ligands, receptors,
and transducers exists but only a few second messengers and types of effector enzymes
are known.

Receptor tyrosine kinases have a simpler mechanism for signal transduction. With
these enzymes, the membrane receptor, transducer, and effector enzyme are combined
in one enzyme. A receptor domain on the extracellular side of the membrane is con-
nected to the cytosolic active site by a transmembrane segment. The active site catalyzes
phosphorylation of its target proteins.

Amplification is an important feature of signaling pathways. A single ligand receptor
complex can interact with a number of transducer molecules, each of which can acti-
vate several molecules of effector enzyme. Similarly, the production of many second
messenger molecules can activate many kinase molecules that catalyze the phosphoryla-
tion of many target proteins. This series of amplification events is called a cascade. The
cascade mechanism means that small amounts of an extracellular compound can affect
large numbers of intracellular enzymes without crossing the plasma membrane or
binding to each target protein.

Not all chemical stimuli follow the general mechanism of signal transduction
shown in Figure 9.40. For example, because steroid hormones are hydrophobic, they
can diffuse across the plasma membrane into the cell where they can bind to specific re-
ceptor proteins in the cytoplasm. The steroid receptor complexes are then transferred to
the nucleus. The complexes bind to specific regions of DNA called hormone response el-
ements and thereby enhance or suppress the expression of adjacent genes.

Kinases were introduced in Section 6.9.

KEY CONCEPT

Membrane receptors are the primary step in
carrying information across a membrane.

The actions of the hormones insulin,
glucagon, and epinephrine and the
roles of transmembrane signaling path-
ways in the regulation of carbohydrate
and lipid metabolism are described in
Sections 11.5, 13.3, 13.7, 13.10,

16. 1C, 16.4 (Box), and 16.7.

B. Signal Transducers

There are many kinds of receptors and many different transducers. Bacterial transduc-
ers are different than eukaryotic ones. There are some eukaryotic transducers found in
most species. In this section, we’ll concentrate on those general transducers.

Many membrane receptors interact with a family of guanine nucleotide binding
proteins called G proteins. G proteins act as transducers — the agents that transmit external

286 CHAPTER 9 Lipids and Membranes

Figure 9.41 ►

Hydrolysis of guanosine 5'-triphosphate (GTP)
to guanosine 5 '-diphosphate (GDP) and phos-
phate (Pj).

Hormone receptor
complex

GDP GTP

▲ Figure 9.42

G-protein cycle. G proteins undergo activation after binding to a
receptor ligand complex and are slowly inactivated by their own
GTPase activity. Both G^-GTP/GDP and G^are membrane-
bound.

stimuli to effector enzymes. G proteins have GTPase activity; that is, they
slowly catalyze hydrolysis of bound guanosine 5 '-triphosphate (GTP, the
guanine analog of ATP) to guanosine 5 '-diphosphate (GDP) (Figure 9.41).
When GTP is bound to G protein it is active in signal tranduction and
when G protein is bound to GDP it is inactive. The cyclic activation and
deactivation of G proteins is shown in Figure 9.42. The G proteins in-
volved in signaling by hormone receptors are peripheral membrane pro-
teins located on the inner surface of the plasma membrane. Each protein
consists of an a, a /3, and a / subunit. The a and /subunits are lipid an-
chored membrane proteins; the a subunit is a fatty acyl anchored pro-
tein and the /subunit is a prenyl anchored protein. The complex of G a py
and GDP is inactive.

When a hormone receptor complex diffusing laterally in the mem-
brane encounters and binds G a p r it induces the G protein to change to
an active conformation. Bound GDP is rapidly exchanged for GTP pro-
moting the dissociation of G^-GTP from Gpy. Activated G^-GTP then
interacts with the effector enzyme. The GTPase activity of the G protein
acts as a built-in timer since G proteins slowly catalyze the hydrolysis of
GTP to GDP. When GTP is hydrolyzed the G^-GDP complex reassoci-
ates with G^and the G^^-GDP complex is regenerated. G proteins
have evolved into good switches because they are very slow catalysts,
typically having a fc cat of only about 3 min -1 .

G proteins are found in dozens of signaling pathways including the
adenylyl cyclase and the inositol-phospholipid pathways discussed
below. An effector enzyme can respond to stimulatory G proteins (Gs)
or inhibitory G proteins (Gi). The a subunits of different G proteins are
distinct providing varying specificity but the /3 and /subunits are similar
and often interchangeable. Humans have two dozen a proteins, five /?
proteins, and six / proteins.

9.11 Transduction of Extracellular Signals 287

C. The Adenylyl Cyclase Signaling Pathway

The cyclic nucleotides 3 ',5 '-cyclic adenosine monophosphate (cAMP) and its guanine
analog, 3 ',5 '-cyclic guanosine monophosphate (cGMP), are second messengers that
help transmit signals from external sources to intracellular enzymes. cAMP is produced
from ATP by the action of adenylyl cyclase (Figure 9.43) and cGMP is formed from
GTP in a similar reaction.

Many hormones that regulate intracellular metabolism exert their effects on target
cells by activating the adenylyl cyclase signaling pathway. Binding of a hormone to a
stimulatory receptor causes the conformation of the receptor to change promoting in-
teraction between the receptor and a stimulatory G protein, G s . The receptor ligand
complex activates G s that, in turn, binds the effector enzyme adenylyl cyclase and acti-
vates it by allosterically inducing a conformational change at its active site.

Adenylyl cyclase is an integral membrane enzyme whose active site faces the cy-
tosol. It catalyzes the formation of cAMP from ATP. cAMP then diffuses from the mem-
brane surface through the cytosol and activates an enzyme known as protein kinase A.
This kinase is made up of a dimeric regulatory subunit and two catalytic subunits and is
inactive in its fully assembled state. When the cytosolic concentration of cAMP in-
creases as a result of signal transduction through adenylyl cyclase, four molecules of
cAMP bind to the regulatory subunit of the kinase releasing the two catalytic subunits,
which are enzymatically active (Figure 9.44). Protein kinase A, a serine-threonine pro-
tein kinase, catalyzes phosphorylation of the hydroxyl groups of specific serine and
threonine residues in target enzymes. Phosphorylation of amino acid side chains on the
target enzymes is reversed by the action of protein phosphatases that catalyze hydrolytic
removal of the phosphoryl groups.

The ability to turn off a signal transduction pathway is an essential element of all
signaling processes. For example, the cAMP concentration in the cytosol increases only
transiently. A soluble cAMP phosphodiesterase catalyzes the hydrolysis of cAMP to
AMP (Figure 9.43) limiting the lifetime of the second messenger. At high concentra-
tions, the methylated purines caffeine and theophylline (Figure 9.45) inhibit cAMP
phosphodiesterase, thereby decreasing the rate of conversion of cAMP to AMR These
inhibitors prolong and intensify the effects of cAMP and hence the activating effects of
the stimulatory hormones.

Hormones that bind to stimulatory receptors activate adenylyl cyclase and raise in-
tracellular cAMP levels. Hormones that bind to inhibitory receptors inhibit adenylyl cy-
clase activity via receptor interaction with the transducer G|. The ultimate response of a
cell to a hormone depends on the type of receptors present and the type of G protein to
which they are coupled. The main features of the adenylyl cyclase signaling pathway, in-
cluding G proteins, are summarized in Figure 9.46.

D. The Inositol-Phospholipid Signaling Pathway

Another major signal transduction pathway produces two different second messengers,
both derived from a plasma membrane phospholipid called phosphatidylinositol 4,5-
Hsphosphate (PIP 2 ) (Figure 9.47). PIP 2 is a minor component of plasma membranes
located in the inner monolayer. It is synthesized from phosphatidylinositol by two suc-
cessive phosphorylation steps catalyzed by ATP- dependent kinases.

Following binding of a ligand to a specific receptor, the signal is transduced
through the G protein G q . The active GTP-bound form of G q activates the effector en-
zyme phosphoinositide-specific phospholipase C that is bound to the cytoplasmic
face of the plasma membrane. Phospholipase C catalyzes the hydrolysis of PIP 2 to in-
ositol 1,4,5-tnsphosphate (IP 3 ) and diacylglycerol (Figure 9.47). Both IP 3 and diacyl-
glycerol are second messengers that transmit the original signal to the interior of
the cell.

IP 3 diffuses through the cytosol and binds to a calcium channel in the membrane
of the endoplasmic reticulum. This causes the calcium channel to open for a short time,
releasing Ca ® from the lumen of the endoplasmic reticulum into the cytosol. Calcium
is also an intracellular messenger because it activates calcium-dependent protein

©.

o— P —

G 0 — P =

e o — P =

©,

H2O-X

H® <r^

\ /

cAMP

phosphodiesterase

▲ Figure 9.43

Production and inactivation of cAMP. ATP is

converted to cAMP by the transmembrane
enzyme adenylyl cyclase. The second mes-
senger is subsequently converted to 5'-AMP
by the action of a cytosolic cAMP phospho-
diesterase.

The response of E. coli to changes in
glucose concentrations, modulated by
cAMP, is described in Section 21. 7B.

288 CHAPTER 9 Lipids and Membranes

R R

Inactive complex

c ^

Active catalytic subunits

▲ Figure 9.44

Activation of protein kinase A. The assembled
complex is inactive. When four molecules of
cAMP bind to the regulatory subunit (R) dimer,
the catalytic subunits (C) are released.

ch 3

Theophylline

▲ Figure 9.45
Caffeine and theophylline.

kinases that catalyze phosphorylation of various protein targets. The calcium signal is
short-lived since Ca^ is pumped back into the lumen of the endoplasmic reticulum
when the channel closes.

The other product of PIP 2 hydrolysis, diacylglycerol, remains in the plasma mem-
brane. Protein kinase C, which exists in equilibrium between a soluble cytosolic form
and a peripheral membrane form, moves to the inner face of the plasma membrane
where it binds transiently and is activated by diacylglycerol and Ca . Protein kinase C
catalyzes phosphorylation of many target proteins altering their catalytic activity.
Several protein kinase C isozymes exist, each with different catalytic properties and
tissue distribution. They are members of the serine-threonine kinase family.

Signaling via the inositol-phospholipid pathway is turned off in several ways. First,
when GTP is hydrolyzed, G q returns to its inactive form and no longer stimulates phos-
pholipase C. The activities of IP 3 and diacylglycerol are also transient. IP 3 is rapidly hy-
drolyzed to other inositol phosphates (which can also be second messengers) and inositol.
Diacylglycerol is rapidly converted to phosphatidate. Both inositol and phosphatidate are
recycled back to phosphatidylinositol. The main features of the inositol-phospholipid
signaling pathway are summarized in Figure 9.48.

Phosphatidylinositol is not the only membrane lipid that gives rise to second mes-
sengers. Some extracellular signals lead to the activation of hydrolases that catalyze the
conversion of membrane sphingolipids to sphingosine, sphingosine 1 -phosphate, or
ceramide. Sphingosine inhibits protein kinase C, and ceramide activates a protein ki-
nase and a protein phosphatase. Sphingosine 1 -phosphate can activate phospholipase

Stimulatory

hormone

Inhibitory

hormone

kinase A
(active)

Protein — OFI

■> Protein — ©

Cellular

response

Figure 9.46 ▲

Summary of the adenylyl cyclase signaling pathway. Binding of a hormone to a stimulatory transmem-
brane receptor (R s ) leads to activation of the stimulatory G protein (G s ) on the inside of the mem-
brane. Other hormones can bind to inhibitory receptors (Rj) that are coupled to adenylyl cyclase by
the inhibitory G protein Gj. G s activates the integral membrane enzyme adenylyl cyclase whereas Gj
inhibits it. cAMP activates protein kinase A resulting in the phosphorylation of cellular proteins.

9.11 Transduction of Extracellular Signals 289

Rt — C — O — CH 2

r 2 — c — o— ch
II I

o ch 2

Phosphatidylinositol 4,5-b/sphosphate
(PIP 2 )

◄ Figure 9.47

Phosphatidylinositol 4,5-Z;/sphosphate (PIP 2 ).

Phosphatidylinositol 4,5-b/'sphosphate
(PIP 2 ) produces two second messengers, in-
ositol l,4,5-f/7sphosphate (IP 3 ) and diacyl-
glycerol. PIP 2 is synthesized by the addition
of two phosphoryl groups (red) to phos-
phatidylinositol and hydrolyzed to IP 3 and
diacylglycerol by the action of a phospho-
inositide-specific phospholipase C.

Phospholipase C

/- H2 °

Diacylglycerol

Rt — C — O — CH 2
R 2 — C— O — CH

II I

O CH 2 — OH

Inositol 1,4,5-tr/sphosphate
(IP3)

D, which specifically catalyzes hydrolysis of phosphatidylcholine. The phosphatidate
and the diacylglycerol formed by this hydrolysis appear to be second messengers. The
full significance of the wide variety of second messengers generated from membrane
lipids (each with its own specific fatty acyl groups) has not yet been determined.

EXTERIOR

Endoplasmic
reticulum

Protein — OH

Protein—®

Cellular

response

Phosphatases

Cellular

response

◄ Figure 9.48

Inositol-phospholipid signaling pathway.

Binding of a ligand to its transmembrane re-
ceptor (R) activates the G protein (G q ). This
in turn stimulates a specific membrane-
bound phospholipase C (PLC) that catalyzes
hydrolysis of the phospholipid PI P 2 in the
inner leaflet of the plasma membrane. The
resulting second messengers, IP 3 and diacyl-
glycerol (DAG), are responsible for carrying
the signal to the interior of the cell. IP 3 dif-
fuses to the endoplasmic reticulum where it
binds to and opens a Ca^ channel in the
membrane releasing stored Ca®. Diacyl-
glycerol remains in the plasma membrane
where it — along with Ca^ — activates the
enzyme protein kinase C (PKC).

290 CHAPTER 9 Lipids and Membranes

BOX 9.7 BACTERIAL TOXINS AND G PROTEINS

G proteins are the biological targets of cholera and pertussis
(whooping cough) toxins that are secreted by the disease-
producing bacteria Vibrio cholerae and Bordetella pertussis ,
respectively. Both diseases involve overproduction of cAMP.

Cholera toxin binds to ganglioside G M1 on the cell surface
(Section 9.5) and a subunit of it crosses the plasma membrane
and enters the cytosol. This subunit catalyzes covalent modifi-
cation of the a subunit of the G protein G s inactivating its GT-
Pase activity. The adenylyl cyclase of these cells remains acti-
vated and cAMP levels stay high. In people infected with
cholerae , cAMP stimulates certain transporters in the plasma
membrane of the intestinal cells leading to a massive secretion
of ions and water into the gut. The dehydration resulting from
diarrhea can be fatal unless fluids are replenished.

Pertussis toxin binds to a glycolipid called lactosylceramide
found on the cell surface of epithelial cells in the lung. It is taken
up by endocytosis. The toxin catalyzes covalent modification of
Gi. In this case, the modified G protein is unable to replace

GDP with GTP and therefore adenylyl cyclase activity cannot
be reduced via inhibitory receptors. The resulting increase in
cAMP levels produces the symptoms of whooping cough.

► Pertussis toxin. The bacterial
toxin has five different subunits
colored red, green, blue, purple,
and yellow. [PDB 1BCP]

Ligands

ligand binding and
dimerization

n ATP-^
n ADP^

autophosphorylation

E. Receptor Tyrosine Kinases

Many growth factors operate by a signaling pathway that includes a multifunctional
transmembrane protein called a receptor tyrosine kinase. As shown in Figure 9.49, the
receptor, transducer, and effector functions are all found in a single membrane protein.
In one type of activation, a ligand binds to the extracellular domain of the receptor,
activating tyrosine kinase catalytic activity in the intracellular domain by dimerization
of the receptor. When two receptor molecules associate, each tyrosine kinase domain
catalyzes the phosphorylation of specific tyrosine residues of its partner, a process called
autophosphorylation. The activated tyrosine kinase then catalyzes phosphorylation of
certain cytosolic proteins, setting off a cascade of events in the cell.

The insulin receptor is an a 2 p 2 tetramer (Figure 9.50). When insulin binds to the
a subunit, it induces a conformational change that brings the tyrosine kinase domains
of the (3 subunits together. Each tyrosine kinase domain in the tetramer catalyzes the
phosphorylation of the other kinase domain. The activated tyrosine kinase also cat-
alyzes the phosphorylation of tyrosine residues in other proteins that help regulate nutrient
utilization.

Recent research has found that many of the signaling actions of insulin are medi-
ated through PIP 2 (Section 9.12C and Figure 9.51). Rather than causing hydrolysis of
PIP 2 , insulin (via proteins called insulin receptor substrates, IRSs) activates phospho-
tidylinositol 3-kinase, an enzyme that catalyzes the phosphorylation of PIP 2 to
phosphatidylinositol 3,4,5-tnsphosphate (PIP 3 ). PIP 3 is a second messenger that tran-
siently activates a series of target proteins, including a specific phosphoinositide-
dependent protein kinase. In this way, phosphotidylinositol 3 -kinase is the molecular
switch that regulates several serine-threonine protein kinase cascades.

◄ Figure 9.49

Activation of receptor tyrosine kinases. Activation occurs as a result of ligand induced receptor
dimerization. Each kinase domain catalyzes phosphorylation of its partner. The phosphorylated
dimer can catalyze phosphorylation of various target proteins.

Summary 291

Insulin

▲ Figure 9.51

Insulin-stimulated formation of phosphatidylinositol 3,4,5-fr/sphosphate (PIP3). Binding of insulin to its
receptor activates the protein tyrosine kinase activity of the receptor leading to the phosphorylation
of insulin receptor substrates (IRSs). The phosphorylated IRSs interact with phosphotidylinositiol
3 -kinase (PI kinase) at the plasma membrane where the enzyme catalyzes the phosphorylation of
PI P 2 to PIP3. PIP3 acts as a second messenger carrying the message from extracellular insulin to
certain intracellular protein kinases.

Insulin

domains

Phosphoryl groups are removed from both the growth factor receptors and their
protein targets by the action of protein tyrosine phosphatases. Although only a few of
these enzymes have been studied, they appear to play an important role in regulating
the tyrosine kinase signaling pathway. One means of regulation appears to be the local-
ized assembly and separation of enzyme complexes.

▲ Figure 9.50

Insulin receptor. Two extracellular a chains,
each with an insulin binding site, are linked
to two transmembrane p chains, each with
a cytosolic tyrosine kinase domain. Following
insulin binding to the a chains, the tyrosine
kinase domain of each p chain catalyzes
autophosphorylation of tyrosine residues in
the adjacent kinase domain. The tyrosine
kinase domains also catalyze the phospho-
rylation of proteins called insulin receptor
substrates (IRSs).

Summary

1. Lipids are a diverse group of water- insoluble organic compounds.

2. Fatty acids are monocarboxylic acids, usually with an even num-
ber of carbon atoms ranging from 12 to 20.

3. Fatty acids are generally stored as triacylglycerols (fats and oils),
which are neutral and nonpolar.

4. Glycerophospholipids have a polar head group and nonpolar
fatty acyl tails linked to a glycerol backbone.

5. Sphingolipids, which occur in plant and animal membranes, con-
tain a sphingosine backbone. The major classes of sphingolipids
are sphingomyelins, cerebrosides, and gangliosides.

6. Steroids are isoprenoids containing four fused rings.

7. Other biologically important lipids are waxes, eicosanoids, lipid
vitamins, and terpenes.

8. The structural basis for all biological membranes is the lipid
bilayer that includes amphipathic lipids such as glycerophospho-
lipids, sphingolipids, and sometimes cholesterol. Lipids can dif-
fuse rapidly within a leaflet of the bilayer.

9. A biological membrane contains proteins embedded in or associated
with a lipid bilayer. The proteins can diffuse laterally within the
membrane.

10 . Most integral membrane proteins span the hydrophobic
interior of the bilayer, but peripheral membrane proteins are
more loosely associated with the membrane surface. Lipid an-
chored membrane proteins are covalently linked to lipids in the
bilayer.

11. Some small or hydrophobic molecules can diffuse across the bi-
layer. Channels, pores, and passive and active transporters medi-
ate the movement of ions and polar molecules across membranes.
Macromolecules can be moved into and out of the cell by endocy-
tosis and exocytosis, respectively.

12. Extracellular chemical stimuli transmit their signals to the cell in-
terior by binding to receptors. A transducer passes the signal to an
effector enzyme, which generates a second messenger. Signal
transduction pathways often include G proteins and protein
kinases. The adenylyl cyclase signaling pathway leads to activation
of the cAMP- dependent protein kinase A. The inositol-phospho-
lipid signaling pathway generates two second messengers and
leads to the activation of protein kinase C and an increase in the
cytosolic Ca© concentration. In receptor tyrosine kinases, the
kinase is part of the receptor protein.

292 CHAPTER 9 Lipids and Membranes

Problems

1. Write the molecular formulas for the following fatty acids:

(a) nervonic acid (ds- A 15 -tetracosenoate; 24 carbons);

(b) vaccenic acid (ds- A n -octadecenoate); and (c) EPA (all
ds- A 5,8,11, 14,17 -eicosapentaenoate).

2. Write the molecular formulas for the following modified fatty
acids:

(a) lO-(Propoxy) decanoate, a synthetic fatty acid with antipara-
sitic activity used to treat African sleeping sickness, a disease
caused by the protozoan T. brucei (the propoxy group is
— O — CH 2 CH 2 CH 3 )

(b) Phytanic acid (3,7,1 1,1 5-tetramethylhexadecanoate), found
in dairy products

3. Fish ois are rich sources of omega-3 and polyunsaturated fatty
acids and omega- 6 fatty acids are relatively abundant in corn and
sunflower oils. Classify the following fatty acids as omega-3,
omega-6, or neither: (a) linolenate, (b) linoleate, (c) arachido-
nate, (d) oleate, (e) A 8,11,14 -eicosatrienoate.

4. Mammalian platelet activating factor (PAF), a messenger in signal
transduction, is a glycerophospholipid with an ether linkage at C-l.
PAF is a potent mediator of allergic responses, inflammation, and the
toxic-shock syndrome. Draw the structure of PAF (l-alkyl-2-acetyl-
phosphatidyl-choline), where the 1 -alkyl group is a C 16 chain.

5. Docosahexaenoic acid, 22:6 A 4 ’ 7 ’ 10 ’ 13,16,19 , is the predominate
fatty acyl group in the C-2 position of glycerol-3-phosphate in
phosphatidylethanolamine and phosphatidylcholine in many
types of fish.

(a) Draw the structure of docosahexaenoic acid (all double
bonds are cis ) .

(b) Classify docosahexaenoic acid as an omega-3, omega-6, or
omega-9 fatty acid.

6. Many snake venoms contain phospholipase A 2 that catalyzes the
degradation of glycerophospholipids into a fatty acid and a
“lysolecithin.” The amphipathic nature of lysolecithins allows them
to act as detergents in disrupting the membrane structure of red
blood cells, causing them to rupture. Draw the structures of phos-
phatidyl serine (PS) and the products (including a lysolecithin) that
result from the reaction of PS with phospholipase A 2 .

7. Draw the structures of the following membrane lipids:

(a) 1 - stearoyl-2 - oleoyl- 3 -phosphatidylethanolamine

(b) palmitoylsphingomyelin

8. (a) The steroid cortisol participates in the control of carbohy-

drate, protein, and lipid metabolism. Cortisol is derived from
cholesterol and possesses the same four-membered fused ring
system but with: (1) a C-3 keto group, (2) C-4-C-5 double
bond (instead of the C-5-C-6 as in cholesterol), (3) a C-ll
hydroxyl, and (4) a hydroxyl group and a — C(0)CH 2 0H
group at C-l 7. Draw the structure of cortisol.

(b) Ouabain is a member of the cardiac glycoside family found in
plants and animals. This steroid inhibits Na©-K© ATPase
and ion transport and may be involved in hypertension and
high blood pressure in humans. Ouabain possesses a four-
membered fused ring system similar to cholesterol but has
the following structural features: (1) no double bonds in the

rings, (2) hydroxy groups on C-l, C-5, C-ll, and C-14,
(3) — CH 2 OH on C-19, (4) 2-3 unsaturated five-membered
lactone ring on C-l 7 (attached to C-3 of lactone ring), and
(5) 6-deoxymannose attached /3 - 1 to the C-3 oxygen. Draw
the structure of ouabain.

9 . A consistent response in many organisms to changing environ-
mental temperatures is the restructuring of cellular membranes.
In some fish, phosphatidylethanolamine (PE) in the liver micro-
somal lipid membrane contains predominantly docosahexaenoic
acid, 22:6 A 4,7,10,13,16,19 at C-2 of the glycerol-3 -phosphate back-
bone and then either a saturated or monounsaturated fatty acyl
group at C-l. The percentage of the PE containing saturated or
monounsaturated fatty acyl groups was determined in fish accli-
mated at 10°C or 30°C. At 10°C, 61% of the PE molecules con-
tained saturated fatty acyl groups at C-l, and 39% of the PE mol-
ecules contained monounsaturated fatty acyl groups at C-l.
When fish were acclimated to 30°C, 86% of the PE lipids con-
tained saturated fatty acyl groups at C-l, while 14% of the PE
molecules had monounsaturated acyl groups at C-l [Brooks, S.,
Clark, G.T., Wright, S.M., Trueman, R.J., Postle, A.D., Cossins,
A.R., and Maclean, N.M. (2002). Electrospray ionisation mass
spectrometric analysis of lipid restructuring in the carp ( Cyprinus
carpio L.) during cold acclimation./. Exp. Biol 205:3989-3997].
Explain the purpose of the membrane restructuring observed
with the change in environmental temperature.

10 . A mutant gene ( ras ) is found in as many as one- third of all
human cancers including lung, colon, and pancreas, and may be
partly responsible for the altered metabolism in tumor cells. The
ras protein coded for by the ras gene is involved in cell signaling
pathways that regulate cell growth and division. Since the ras pro-
tein must be converted to a lipid anchored membrane protein in
order to have cell- signaling activity, the enzyme farnesyl trans-
ferase (FT) has been selected as a potential chemotherapy target
for inhibition. Suggest why FT might be a reasonable target.

11. Glucose enters some cells by simple diffusion through channels or
pores, but glucose enters red blood cells by passive transport. On
the plot below, indicate which line represents diffusion through a
channel or pore and which represents passive transport. Why do
the rates of the two processes differ?

Extracellular glucose concentration

12. The pH gradient between the stomach (pH 0. 8-1.0) and the gas-
tric mucosal cells lining the stomach (pH 7.4) is maintained by an
H©-K© ATPase transport system that is similar to the ATP-
driven Na©-K© ATPase transport system (Figure 9.38). The
H©-K© ATPase antiport system uses the energy of ATP to pump
H© out of the mucosal cells (me) into the stomach (st) in ex-
change for K© ions. The K© ions that are transported into the
mucosal cells are then cotransported back into the stomach along

Selected Readings 293

with Cl® ions. The net transport is the movement of HC1 into
the stomach.

K© (mc) + Cl© (mc) + H© (mc) + K© (st) + ATP

K© (st) + Cl© (st) + H© (st) + K© (mc) + ADP + P;

Draw a diagram of this H®-K® ATPase system.

13. Chocolate contains the compound theobromine, which is struc-
turally related to caffeine and theophylline. Chocolate products
may be toxic or lethal to dogs because these animals metabolize
theobromine more slowly than humans. The heart, central nerv-
ous system, and kidneys are affected. Early signs of theobromine
poisoning in dogs include nausea and vomiting, restlessness, diar-
rhea, muscle tremors, and increased urination or incontinence.
Comment on the mechanism of toxicity of theobromine in dogs.

ch 3

Theobromine

14. In the inositol signaling pathway, both IP 3 and diacylglycerol
(DAG) are hormonal second messengers. If certain protein ki-
nases in cells are activated by binding Ca©, how do IP 3 and DAG
act in a complementary fashion to elicit cellular responses inside
cells?

15. In some forms of diabetes, a mutation in the (3 subunit of the in-
sulin receptor abolishes the enzymatic activity of that subunit.
How does the mutation affect the cell’s response to insulin? Can
additional insulin (e.g., from injections) overcome the defect?

16. The ras protein (described in Problem 10) is a mutated G protein
that lacks GTPase activity. How does the absence of this activity
affect the adenylyl cyclase signaling pathway?

17. At the momentof fertilization a female egg is about 100/im in di-
ameter. Assuming that each lipid molecule in the plasma mem-
brane has a suface area of 10 -14 cm 2 , how many lipid molecules
are there in the egg plasma membrane if 25% of the surface is
protein?

18. Each fertilized egg cell (zygote) divides 30 times to produce all the
eggs that a female child will need in her lifetime. One of these eggs
will be fertilized giving rise to a new generation. If lipid molecles
are never degraded, how many lipid molecules have you inherited
that were synthesized in your grandmother?

Selected Readings

General

Gurr, M. I., and Harwood, J. L. (1991). Lipid Bio-
chemistry: An Introduction , 4th ed. (London:
Chapman and Hall).

Lester, D. R., Ross, J. J., Davies, P. J., and Reid, J. B.
(1997). Mendels stem length gene ( Le ) encodes a
gibberellin 3 beta-hydroxylase. Plant Cell.
9:1435-1443.

Vance, D. E., and Vance, J. E., eds. (2008).
Biochemistry of Lipids, Lipoproteins, and
Membranes, 5th ed. (New York: Elsevier).

Membranes

Dowhan, W. (1997). Molecular basis for
membrane phospholipid diversity: why are
there so many lipids? Annu. Rev. Biochem.
66:199-232.

lacobson, K., Sheets, E. D., and Simson, R. (1995).
Revisiting the fluid mosaic model of membranes.
Science 268:1441-1442.

Koga, Y., and Morii, H. (2007). Biosynthesis of
ether- type polar lipids in Archaea and evolution-
ary considerations. Microbiol, and Molec. Biol. Rev.
71: 97-120.

Lai, E.C. (2003) Lipid rafts make for slippery plat-
forms./. Cell Biol. 162:365-370.

Lingwood, D., and Simons, K. (2010). Lipid rafts
as a membrane -organizing principle. Science.
327:46-50.

Simons, K., and Ikonen, E. (1997). Lunctional rafts
in cell membranes. Nature. 387:569-572.

Singer, S. J. (1992). The structure and function of
membranes: a personal memoir. /. Membr. Biol.
129:3-12.

Singer, S. J. (2004) Some early history of membrane
molecular biology. Annu. Rev. Physiol. 66:1-27.

Singer, S. J., and Nicholson, G. L. (1972). The fluid
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Introduction
to Metabolism

I n the preceding chapters, we described the structures and functions of the major
components of living cells from small molecules to polymers to larger aggregates
such as membranes. The next nine chapters focus on the biochemical activities that
assimilate, transform, synthesize, and degrade many of the nutrients and cellular com-
ponents already described. The biosynthesis of proteins and nucleic acids, which represent
a significant proportion of the activity of all cells, will be described in Chapters 20-22.

We now move from molecular structure to the dynamics of cell function. Despite
the marked shift in our discussion, we will see that metabolic pathways are governed by
basic chemical and physical laws. By taking a stepwise approach that builds on the foun-
dations established in the first two parts of this book, we can describe how metabolism
operates. In this chapter, we discuss some general themes of metabolism and the ther-
modynamic principles that underlie cellular activities.

For most metabolic sequences neither
the substrate concentration nor the
product concentration changes
significantly ; even though the flux
through the pathway may change
dramatically

— Jeremy R. Knowles (1 989)

10.1 Metabolism Is a Network of Reactions

Metabolism is the entire network of chemical reactions carried out by living cells.
Metabolites are the small molecules that are intermediates in the degradation or biosyn-
thesis of biopolymers. The term intermediary metabolism is applied to the reactions
involving these low- molecular- weight molecules. It is convenient to distinguish between
reactions that synthesize molecules (anabolic reactions) and reactions that degrade
molecules (catabolic reactions).

Anabolic reactions are those responsible for the synthesis of all compounds needed
for cell maintenance, growth, and reproduction. These biosynthesis reactions make
simple metabolites such as amino acids, carbohydrates, coenzymes, nucleotides, and

Top: The fundamental principles of metabolism are the same in animals and plants and in all other organisms.

294

10.1 Metabolism Is a Network of Reactions 295

Light

(photosynthetic
organisms only)

molecules

◄ Figure 10.1

Anabolism and catabolism. Anabolic
reactions use small molecules and chemical
energy in the synthesis of macromolecules
and in the performance of cellular work.
Solar energy is an important source of meta-
bolic energy in photosynthetic bacteria and
plants. Some molecules, including those
obtained from food, are catabolized to release
energy and either monomeric building
blocks or waste products.

fatty acids. They also produce larger molecules such as proteins, polysaccharides,
nucleic acids, and complex lipids (Figure 10.1).

In some species, all of the complex molecules that make up a cell are synthesized from
inorganic precursors (carbon dioxide, ammonia, inorganic phosphates, etc.) (Section 10.3).
Some species derive energy from these inorganic molecules or from the creation of
membrane potential (Section 9.11). Photosynthetic organisms use light energy to drive
biosynthesis reactions (Chapter 15).

Catabolic reactions degrade large molecules to liberate smaller molecules and
energy. All cells carry out degradation reactions as part of their normal cell metabolism
but some species rely on them as their only source of energy. Animals, for example, re-
quire organic molecules as food. The study of these energy-producing catabolic reactions
in mammals is called fuel metabolism. The ultimate source of these fuels is a biosyn-
thetic pathway in another species. Keep in mind that all catabolic reactions involve the
breakdown of compounds that were synthesized by a living cell — either the same cell, a
different cell in the same individual, or a cell in a different organism.

There is a third class of reactions called amphibolic reactions. They are involved in
both anabolic and catabolic pathways.

Whether we observe bacteria or large multicellular organisms, we find a bewilder-
ing variety of biological adaptations. More than 10 million species may be living on
Earth and several hundred million species may have come and gone throughout the
course of evolution. Multicellular organisms have a striking specialization of cell types
or tissues. Despite this extraordinary diversity of species and cell types the biochemistry of
living cells is surprisingly similar not only in the chemical composition and structure of
cellular components but also in the metabolic routes by which the components are
modified. These universal pathways are the key to understanding metabolism. Once
you’ve learned about the fundamental conserved pathways you can appreciate the addi-
tional pathways that have evolved in some species.

The complete sequences of the genomes of a number of species have been determined.
For the first time we are beginning to have a complete picture of the entire metabolic
network of these species based on the sequences of the genes that encode metabolic enzymes.
Escherichia coli , for example, has about 900 genes that encode enzymes used in interme-
diary metabolism and these enzymes combine to create about 130 different pathways.

KEY CONCEPT

Most of the fundamental metabolic
pathways are present in all species.

296 CHAPTER 10 Introduction to Metabolism

Figure 10.2 ►

A protein interaction network for yeast
( Saccharomyces cerevisiae). Dots represent
individual proteins, colored according to
function. Solid lines represent interactions
between proteins. The colored clusters
identify the large number of genes involved
in metabolism.

Peroxisome

Secretion &

Jding

lation

Mitochondria

Metabolism &
amino acid
biosynthesis

RNA

processing

vesicle
\.'f\ . transport

Chromatin & <
transcription

cytoplasmic

transport

Nuclear
migration
& protein
degradation

Cell wall
biosynthesis

Cell polarity &
morphogenesis

Mitosis & chr.
segregation

DNA replication
& repair

These metabolic genes account for 21% of the genes in the genome. Other species of
bacteria have a similar number of enzymes that carry out the basic metabolic reactions.
Some species contain additional pathways. The bacterium that causes tuberculosis,
Mycobacterium tuberculosis , has about 250 enzymes involved in fatty acid metabolism —
five times as many as E. coli.

The yeast Saccharomyces cerevisiae is a single-celled member of the fungus king-
dom. Its genome contains 5900 protein-encoding genes. Of these, 1200 (20%) encode
enzymes involved in intermediary and energy metabolism (Figure 10.2). The nematode
Caenorhabditis elegans is a small, multicellular animal with many of the same special-
ized cells and tissues found in larger animals. Its genome encodes 19,100 proteins of
which 5300 (28%) are thought to be required in various pathways of intermediary
metabolism. In the fruit fly, Drosophila melanogaster , approximately 2400 (17%) of its
14,100 genes are predicted to be involved in intermediary metabolic pathways and
bioenergetics. The exact number of genes required for basic metabolism in humans is
not known but it’s likely that about 5000 genes are needed. (The human genome has
approximately 22,000 genes.)

There are five common themes in metabolism.

1. Organisms or cells maintain specific internal concentrations of inorganic ions,
metabolites, and enzymes. Cell membranes provide the physical barrier that segre-
gates cell components from the environment.

2. Organisms extract energy from external sources to drive energy- consuming reac-
tions. Photosynthetic organisms derive energy from the conversion of solar energy
to chemical energy. Other organisms obtain energy from the ingestion and catabolism
of energy-yielding compounds.

3. The metabolic pathways in each organism are specified by the genes it contains in
its genome.

4. Organisms and cells interact with their environment. The activities of cells must be
geared to the availability of energy, organisms grow and reproduce. When the sup-
ply of energy from the environment is plentiful. When the supply of energy from
the environment is limited, energy demands can be temporarily met by using inter-
nal stores or by slowing metabolic rates as in hibernation, sporulation, or seed for-
mation. If the shortage is prolonged, organisms die.

5. The cells of organisms are not static assemblies of mtneylecules. Many cell compo-
nents are continually synthesized and degraded, that is, they undergo turnover , even

10.2 Metabolic Pathways

297

though their concentrations may remain virtually constant. The concentrations of

other compounds change in response to changes in external or internal conditions.

The metabolism section of this book describes metabolic reactions that operate in
most species. For example, enzymes of glycolysis (the degradation of sugar) and of glu-
coneogenesis (biosynthesis of glucose) are present in almost all species. Although most
cells possess the same set of central metabolic reactions, cell and organism differentiation
is possible because of additional enzymatic reactions specific to the tissue or species.

10.2 Metabolic Pathways

The vast majority of metabolic reactions are catalyzed by enzymes so a complete
description of metabolism includes not only the reactants, intermediates, and products of
cellular reactions but also the characteristics of the relevant enzymes. Most cells can per-
form hundreds to thousands of reactions. We can deal with this complexity by systemat-
ically subdividing metabolism into segments or branches. In the following chapters, we
begin by considering separately the metabolism of the four major groups of biomole-
cules: carbohydrates, lipids, amino acids, and nucleotides. Within each of the four areas
of metabolism, we recognize distinct sequences of metabolic reactions, called pathways.

A. Pathways Are Sequences of Reactions

A metabolic pathway is the biological equivalent of a synthesis scheme in organic chem-
istry. A metabolic pathway is a series of reactions where the product of one reaction
becomes the substrate for the next reaction. Some metabolic pathways may consist of
only two steps while others may be a dozen steps in length.

Its not easy to define the limits of a metabolic pathway. In the laboratory, a chemi-
cal synthesis has an obvious beginning substrate and an obvious end product but cellu-
lar pathways are interconnected in ways that make it difficult to pick a beginning and an
end. For example, in the catabolism of glucose (Chapter 11), where does glycolysis
begin and end? Does it begin with polysaccharides (such as glycogen and starch), extra-
cellular glucose, glucose 6-phosphate, or intracellular glucose? Does the pathway end
with pyruvate, acetyl CoA, lactate, or ethanol? Start and end points can be assigned
somewhat arbitrarily, often according to tradition or for ease of study, but keep in mind
that reactions and pathways can be linked to form extended metabolic routes. This net-
work is very obvious when you examine the large metabolic charts that are sometimes
posted on the walls outside professors’ offices (Figure 10.3).

Individual metabolic pathways can take different forms. A linear metabolic pathway,
such as the biosynthesis of serine, is a series of independent enzyme -catalyzed reactions

◄ Figure 10.3

Part of a large metabolic chart published by
Roche Applied Science.

298 CHAPTER 10 Introduction to Metabolism

(a)

3-Phosphoglycerate

3-Phosphohydroxypyruvate

3-Phosphoserine

Serine

▲ Figure 10.4

Forms of metabolic pathways, (a) The biosyn-
thesis of serine is an example of a linear
metabolic pathway. The product of each step
is the substrate for the next step, (b) The se-
quence of reactions in a cyclic pathway
forms a closed loop. In the citric acid cycle,
an acetyl group is metabolized via reactions
that regenerate the intermediates of the
cycle, (c) In fatty acid biosynthesis, a spiral
pathway, the same set of enzymes catalyzes
a progressive lengthening of the acyl chain.

KEY CONCEPT

The limitations of chemistry and physics
dictate that metabolic pathways consist
of many small steps.

in which the product of one reaction is the substrate for the next reaction in the pathway
(Figure 10.4a). A cyclic metabolic pathway, such as the citric acid cycle, is also a sequence
of enzyme -catalyzed steps, but the sequence forms a closed loop, so the intermediates
are regenerated with every turn of the cycle (Figure 10.4b). In a spiral metabolic pathway,
such as the biosynthesis of fatty acids (Section 16.6), the same set of enzymes is used re-
peatedly for lengthening or shortening a given molecule (Figure 10.4c).

Each type of pathway may have branch points where metabolites enter or leave. In
most cases, we don’t emphasize the branching nature of pathways because we want to
focus on the main routes followed by the most important metabolites. We also want to
focus on the pathways that are commonly found in all species. These are the most fun-
damental pathways. Don’t be misled by this simplification. A quick glance at any metabolic
chart will show that pathways have many branch points and that initial substrates and final
products are often intermediates in other pathways. The serine pathway in Figure 10.3 is
a good example. Can you find it?

B. Metabolism Proceeds by Discrete Steps

Intracellular environments don’t change very much. Reactions proceed at moderate
temperatures and pressures, at rather low reactant concentrations, and at close to neu-
tral pH. We often refer to this as homeostasis at the cellular level.

These conditions require a multitude of efficient enzymatic catalysts. Why are so
many distinct reactions carried out in living cells? In principle, it should be possible to
carry out the degradation and the synthesis of complex organic molecules with far
fewer reactions.

One reason for multistep pathways is the limited reaction specificity of enzymes.
Each active site catalyzes only a single step of a pathway. The synthesis of a molecule —
or its degradation — therefore follows a metabolic route defined by the availability of
suitable enzymes. As a general rule, a single enzyme -catalyzed reaction can only break
or form a few covalent bonds at a time. Often the reaction involves the transfer of a sin-
gle chemical group. Thus, the large number of reactions and enzymes is due, in part, to
the limitations of enzymes and chemistry.

Another reason for multiple steps in metabolic pathways is to control energy input
and output. Energy flow is mediated by energy donors and acceptors that carry discrete
quanta of energy. As we will see, the energy transferred in a single reaction seldom
exceeds 60 kj mol -1 . Pathways for the biosynthesis of molecules require the transfer of
energy at multiple points. Each energy- requiring reaction corresponds to a single step
in the reaction sequence.

The synthesis of glucose from carbon dioxide and water requires the input of
-2900 kj mol -1 of energy. It is not thermodynamically possible to synthesize glucose in
a single step (Figure 10.5). Similarly, much of the energy released during a catabolic
process (such as the oxidation of glucose to carbon dioxide and water, which releases
the same 2900 kj mol -1 ) is transferred to individual acceptors one step at a time rather

10.2 Metabolic Pathways

299

(a) Glucose + 6 0 2

V. J

(b)

Glucose + 6 0 2

Impossible

one-step

synthesis

Energy

Multistep

pathway

Energy

6 C0 2 + 6 H 2 0

Anabolism

(Biosynthesis)

Uncontrolled

combustion

Energy - ^ « Energy

Multistep

pathway

^ Energy

v v

^ Energy

6 C0 2 + 6 H 2 0
Catabolism

◄ Figure 10.5

Single-step versus multistep pathways, (a) The

synthesis of glucose cannot be accomplished
in a single step. Multistep synthesis is cou-
pled to the input of small quanta of energy
from ATP and NADH. (b) The uncontrolled
combustion of glucose releases a large
amount of energy all at once. A multistep
enzyme-catalyzed pathway releases the
same amount of energy but conserves much
of it in a manageable form.

than being released in one grand, inefficient explosion. The efficiency of energy transfer
at each step is never 100%, but a considerable percentage of the energy is conserved in
manageable form. Energy carriers that accept and donate energy, such as adenine
nucleotides (ATP) and nicotinamide coenzymes (NADH), are found in all life forms.

A major goal of learning about metabolism is to understand how these “quanta” of
energy are used. ATP and NADH — and other coenzymes — are the “currency” of metab-
olism. This is why metabolism and bioenergetics are so closely linked.

C. Metabolic Pathways Are Regulated

Metabolism is highly regulated. Organisms react to changing environmental conditions
such as the availability of energy or nutrients. Organisms also respond to genetically
programmed instructions. For example, during embryogenesis or reproduction, the
metabolism of individual cells can change dramatically.

The responses of organisms to changing conditions range from small changes to
drastically reorganizing the metabolic processes that govern the synthesis or degrada-
tion of biomolecules and the generation or consumption of energy. Control processes
can affect many pathways or only a few, and the response time can range from less than
a second to hours or longer. The most rapid biological responses, occurring in millisec-
onds, include changes in the passage of small ions (e.g., Na®, K®, and Ca©) through
cell membranes. Transmission of nerve impulses and muscle contraction depend on ion
movement. The most rapid responses are also the most short-lived; slower responses
usually last longer.

It is important to understand some basic concepts of pathways in order to see how
they are regulated. Consider a simple linear pathway that begins with substrate A and
ends with product P.

( 10 . 1 )

300

CHAPTER 10 Introduction to Metabolism

The precise technical term for the con-
dition where cellular pathways are not
in a dynamic steady-state condition
is . . . dead.

Each of the reactions is catalyzed by an enzyme and they are all reversible. Most reac-
tions in living cells have reached equilibrium so the concentrations of B, C, D, and E do
not change very much. This is similar to the steady state condition we encountered in
Section 5.3A. The steady state condition can be visualized by imagining a series of
beakers of different sizes (Figure 10.6). Water flows into the first beaker from a tap and
when it fills up the water spills over into another beaker. After filling up a series of
beakers, there will be a steady flow of water from the tap onto the floor. The rate of flow
is analogous to the flux through a metabolic pathway. The flux can vary from a trickle to
a gusher but the steady state levels of water in each beaker don’t change. (Unfortunately,
this analogy doesn’t allow us to see that in a metabolic pathway the flux could also be in
the opposite direction.)

Flux through a metabolic pathway will decrease if the concentration of the initial
substrate falls below a certain threshold. It will also decrease if the concentration of the
final product rises. These are changes that affect all pathways. However, in addition to
these normal concentration effects, there are special regulatory controls that affect the
activity of particular enzymes in the pathway. It is tempting to visualize regulation of a
pathway by the efficient manipulation of a single rate limiting enzymatic reaction,
sometimes likened to the narrow part of an hourglass. In many cases, however, this is an
oversimplification. Flux through most pathways depends on controls at several steps.
These steps are special reactions in the pathways where the steady state concentrations
of substrates and products are far from the equilibrium concentrations so the flux tends
to go only in one direction. A regulatory enzyme contributes a particular degree of con-
trol over the overall flux of the pathway in which it participates. Because intermediates
or cosubstrates from several sources can feed into or out of a pathway, the existence of
multiple control points is normal; an isolated, linear, pathway is rare.

There are two common patterns of metabolic regulation: feedback inhibition and
feed-forward activation. Feedback inhibition occurs when a product (usually the end
product) of a pathway controls the rate of its own synthesis through inhibition of an
early step, usually the first committed step (the first reaction that is unique to the pathway).

-> B

-> C

-> E

-> P

( 10 . 2 )

The advantage of such a regulatory pattern in a biosynthetic pathway is obvious. When
the concentration of P rises above its steady state level, the effect is transmitted back
through the pathway and the concentrations of each intermediate also rise. This causes
flux to reverse in the pathway, leading to a net increase in the production of product A
from reactant P. Flux in the normal direction is restored when P is depleted. The path-
way is inhibited at an early step; otherwise, metabolic intermediates would accumulate
unnecessarily. The important point in Reaction 10.2 is that the reaction catalyzed by
enzyme El is not allowed to reach equilibrium. It is a metabolically irreversible reaction
because the enzyme is regulated. Flux through this point is not allowed to go in the op-
posite direction.

Feed-forward activation occurs when a metabolite produced early in a pathway acti-
vates an enzyme that catalyzes a reaction further down the pathway.

-> B

-> C

-> D

-> P

(10.3)

▲ Figure 10.6

Steady state and flux in a metabolic pathway.

The rate of flow is equivalent to the flux in a
pathway, and the constant amount of water
in each beaker is analogous to the steady
state concentrations of metabolites in a
pathway.

In this example, the activity of enzyme E : (which converts A to B) is coordinated with
the activity of enzyme E 4 (which converts D to E). An increase in the concentration of
metabolite B increases flux through the pathway by activating E4. (E4 would normally
be inactive in low concentrations of B.)

In Section 5.10, we discussed the modulation of individual regulatory enzymes.
Allosteric activators and inhibitors, which are usually metabolites, can rapidly alter the

10.2 Metabolic Pathways 301

activity of many of these enzymes by inducing conformational changes that affect cat-
alytic activity. We will see many examples of allosteric modulation in the coming chapters.
The allosteric modulation of regulatory enzymes is fast but not as rapid in cells as it can
be with isolated enzymes.

The activity of interconvertible enzymes can also be rapidly and reversibly altered
by covalent modification, commonly by the addition and removal of phosphoryl
groups as described in Section 5.9D. Recall that phosphorylation, catalyzed by protein
kinases at the expense of ATP, is reversed by the action of protein phosphatases, which
catalyze the hydrolytic removal of phosphoryl groups. Individual enzymes differ in
whether their response to phosphorylation is activation or deactivation. Interconvert-
ible enzymes in catabolic pathways are generally activated by phosphorylation and de-
activated by dephosphorylation; most interconvertible enzymes in anabolic pathways
are inactivated by phosphorylation and reactivated by dephosphorylation. The activa-
tion of kinases with multiple specificities allows coordinated regulation of more than
one metabolic pathway by one signal. The cascade nature of intracellular signaling
pathways, described in Section 9.12, also means that the initial signal is amplified
(Figure 10.7).

The amounts of specific enzymes can be altered by increasing the rates of specific
protein synthesis or degradation. This is usually a slow process relative to allosteric or
covalent activation and inhibition. However, the turnover of certain enzymes may be
rapid. Keep in mind that several modes of regulation can operate simultaneously within
a metabolic pathway.

In Part 4 of this book, we examine
more closely the regulation of gene
expression and protein synthesis.

D. Evolution of Metabolic Pathways

The evolution of metabolic pathways is an active area of biochemical research. These
studies have been greatly facilitated by the publication of hundreds of complete genome
sequences, especially prokaryotic genomes. Biochemists can now compare pathway
enzymes in a number of species that show a diverse variety of pathways. Many of these
pathways provide clues to the organization and structure of the primitive pathways that
were present in the first cells.

There are many possible routes to the formation of a new metabolic pathway. The
simplest case is the addition of a new terminal step to a preexisting pathway. Consider
the hypothetical pathway in Equation 10.1. The original pathway might have termi-
nated with the production of metabolite E after a four- step transformation from sub-
strate A. The availability of substantial quantities of metabolite E might favor the evolution
of a new enzyme (E 5 in this case) that could use E as a substrate to make R The pathways

HO— ( ^Protein)

Initial signal

Signal

transduction

ATP

ADP^i

Q— (Protein)

Cellular

response

Protein

kinase

response

Protein

-OH

^-ATP

Sadp

Protein

-G

v 7

Cellular

response

◄ Figure 10.7

Regulatory role of a protein kinase. The effect
of the initial signal is amplified by the sig-
naling cascade. Phosphorylation of different
cellular proteins by the activated kinase
results in coordinated regulation of different
metabolic pathways. Some pathways may
be activated, whereas others are inhibited.

— Q) represents a protein-bound phosphate
group.

302

CHAPTER 10 Introduction to Metabolism

leading to synthesis of asparagine and glutamine from aspartate and glutamate path-
ways are examples of this type of pathway evolution. This forward evolution is thought
to be a common mechanism of evolution of new pathways.

In other cases, a new pathway can form by evolving a branch to a preexisting path-
way. For example, consider the conversion of C to D in the Equation 10.1 pathway. This
reaction is catalyzed by enzyme E 3 . The primitive E 3 enzyme might not have been as
specific as the modern enzyme. In addition to producing product D, it might have syn-
thesized a smaller amount of another metabolite, X. The availability of product X might
have conferred some selective advantage to the cell favoring a duplication of the E 3
gene. Subsequent divergence of the two copies of the gene gave rise to two related
enzymes that specifically catalyzed C — > D and C — > X. There are many examples of
evolution by gene duplication and divergence (e.g., lactate dehydrogenase and malate
dehydrogenase, Section 4.7). (We have mostly emphasized the extreme specificity of
enzyme reactions but, in fact, many enzymes can catalyze several different reactions
using structurally similar substrates and products.)

Some pathways might have evolved “backwards.” A primitive pathway might have
utilized an abundant supply of metabolite E in the environment in order to make prod-
uct P. As the supply of E became depleted over time there was selective pressure to
evolve a new enzyme (E 4 ) that could make use of metabolite D to replenish metabolite
E. When D became rate limiting, cells could gain a selective advantage by utilizing C to
make more metabolite D. In this way the complete modern pathway evolved by
retroevolution , successively adding simpler precursors and extending the pathway.

Sometimes an entire pathway can be duplicated and subsequent adaptive evolution
leads to two independent pathways with homologous enzymes that catalyze related re-
actions. There is good evidence that the pathways leading to biosynthesis of tryptophan
and histidine evolved in this manner. Enzymes can also be recruited from one pathway
for use in another without necessarily duplicating an entire pathway. We’ll encounter
several examples of homologous enzymes that are used in different pathways.

Finally, a new pathway can evolve by “reversing” an existing pathway. In most cases,
there is one step in a pathway that is essentially irreversible. Let’s assume that the third
step in our hypothetical pathway (C — > D) is unable to catalyze the conversion of D to C
because the normal reaction is far from equilibrium. The evolution of a new enzyme
that can catalyze D — > C would allow this entire pathway to reverse direction, converting
P to A. This is how the glycolysis pathway evolved from the glucose biosynthesis (gluco-
neogenesis) pathway. There are many other examples of evolution by pathway reversal.

All of these possibilities play a role in the evolution of new pathways. Sometimes a
new pathway evolves by a combination of different mechanisms of adaptive evolution.
The evolution of the citric acid cycle pathway, which took place several billion years ago,
is an example (Section 12.9). New metabolic pathways are evolving all the time in
response to pesticides, herbicides, antibiotics, and industrial waste. Organisms that can
metabolize these compounds, thus escaping their toxic effects, have evolved new path-
ways and enzymes by modifying existing ones.

10.3 Major Pathways in Cells

This section provides an overview of the organization and function of some central
metabolic pathways that are discussed in subsequent chapters. We begin with the ana-
bolic, or biosynthetic, pathways since these pathways are the most important for growth
and reproduction. A general outline of biosynthetic pathways is shown in Figure 10.8. All
cells require an external source of carbon, hydrogen, oxygen, nitrogen, phosphorus, and
sulfur plus additional inorganic ions (Section 1.2). Some species, notably bacteria and
plants, can grow and reproduce by utilizing inorganic sources of these essential elements.
These species are called autotrophs. There are two distinct categories of autotrophic
species. Heterotrophs, such as animals, need an organic carbon source (e.g., glucose).

Biosynthetic pathways require energy. The most complex organisms (from a bio-
chemical perspective!) can generate useful metabolic energy from sunlight or by oxidiz-
ing inorganic molecules such as NH 4 ®, H 2 , or H 2 S. The energy from these reactions is

10.3 Major Pathways in Cells 303

DNA

RNA

DNA (20)
RNA (21)

Nucleotides

Other

carbohydrates

Pentose phosphate

pathway (1 2.5) G I UCOSe

Starch

Glycogen

Starch synthesis (1 5.5) Light
Glycogen synthesis (12.5)

Ribose,

deoxyribose

Nucleotide
synthesis

Amino

acids

nh 4 @

Calvin
cycle (15.4)

co 2

Gluconeogenesis (12.1)

Photosynthesis (15)

ATP ADP + Pj
NADPH NADP® +

H ©

’yruvate

Fatty acid (16)

o ,n the

a + ir a synthesis (16.1) Fatty
Acetyl CoA acids L| P IC * S

Membranes

Amino acid Protein
synthesis (17) synthesis (22)

— > Amino > Proteins

acids

Nitrogen
N 2 ,NH 4 © fixation (17.1)

used to synthesize the energy-rich compound ATP and the reducing power of NADH.
These cofactors transfer their energy to biosynthetic reactions.

There are two types of autotrophic species. Photoautotrophs obtain most of their en-
ergy by photosynthesis and their main source of carbon is C0 2 . This category
includes photosynthetic bacteria, algae, and plants. Chemoautotrophs obtain their energy
by oxidizing inorganic molecules and utilizing C0 2 as a carbon source. Some bacterial
species are chemoautotrophs but there are no eukaryotic examples.

Heterotrophs can be split into two categories. Photoheterotrophs are photosynthetic
organisms that require an organic compound as a carbon source. There are several
groups of bacteria that are capable of capturing light energy but must rely on some
organic molecules as a carbon source. Chemoheterotrophs are nonphotosynthetic organ-
isms that require organic molecules as carbon sources. Their metabolic energy is usually
derived from the breakdown of the imported organic molecules. We are chemo-
heterotrophs, as are all animals, most protists, all fungi, and many bacteria.

The main catabolic pathways are shown in Figure 10.9. As a general rule, these
degradative pathways are not simply the reverse of biosynthesis pathways. Note that the
citric acid cycle is a major pathway in both anabolic and catabolic metabolism. The
main roles of catabolism are to eliminate unwanted molecules and to generate energy
for use in other processes.

We will examine metabolism in the next few chapters. Our discussion of metabolic
pathways begins in Chapter 1 1 with glycolysis, a ubiquitous pathway for glucose catabolism.
There is a long-standing tradition in biochemistry of introducing students to glycolysis
before any other pathways are encountered. We know a great deal about the reactions in
this pathway and they will illustrate many of the fundamental principles of biochem-
istry. In glycolysis, the hexose is split into two three-carbon metabolites. This pathway
can generate ATP in a process called substrate level phosphorylation. Often, the product
of glycolysis is pyruvate, which can be converted to acetyl CoA for further oxidation.

Chapter 12 describes the synthesis of glucose, or gluconeogenesis. This chapter also
covers starch and glycogen metabolism and outlines the pathway by which glucose is
oxidized to produce NADPH for biosynthetic pathways and ribose for the synthesis of
nucleotides.

The citric acid cycle (Chapter 13) facilitates complete oxidation of the acetate carbons
of acetyl CoA to carbon dioxide. The energy released from this oxidation is conserved in

◄ Figure 10.8

Overview of anabolic pathways. Large mole-
cules are synthesized from smaller ones by
adding carbon (usually in the form of C0 2 )
and nitrogen (usually as NH 4 ®). The main
pathways include the citric acid cycle,
which supplies the intermediates in amino
acid biosynthesis, and gluconeogenesis, which
results in the production of glucose. The
energy for biosynthetic pathways is supplied
by light in photosynthetic organisms or by the
breakdown of inorganic molecules in other
autotrophs. (Numbers in parentheses refer
to the chapters and sections of this book.)

▲ Chemoautotrophs in Yellowstone National
Park. There are many species of Thiobacillus
that derive their energy from the oxidation of
iron or sulfur. They do not require any organic
molecules. The orange and yellow colors
surrounding this hot spring in Yellowstone
National Park are due to the presence of
Thiobacillus. See Chapter 14 for an expla-
nation of how such organisms generate en-
ergy from inorganic molecules.

304 CHAPTER 10 Introduction to Metabolism

Figure 10.9 ►

Overview of catabolic pathways. Amino acids,
nucleotides, monosaccharides, and fatty
acids are formed by enzymatic hydrolysis
of their respective polymers. They are then
degraded in oxidative reactions and energy
is conserved in ATP and reduced coenzymes
(mostly NADH). (Numbers in parentheses RNA

refer to the chapters and sections of this DNA

book.)

Nucleases (20)

Other

carbohydrates

Pentose
phosphate
pathway
(12.5)

Ribose
deoxyribose

Starch

Glycogen

Starch degradation (15.5)
Glycogen degradation (12.5)

Glucose

Nucleotides

Glycolysis (1 1)

Pyruvate

Pyrimidine i (16)

v catabolism (18.9) I „ » . , .. ,* r -,\r ^

a * i a j3-Oxidation(16.7)Fatty^ . . ..
Acetyl CoA< acids ^7 Ll P lds

Purine

catabolism

(18.8)

Uric acid,

urea, NH 4 ©

ATP<.

adp + P j y

Amino acid

degradation (17) Amino Proteases (6.8) _

' T acids < Proteins

nh 3

Electron

transport

NADH

the formation of NADH and ATP. As mentioned above, the citric acid cycle is an essen-
tial part of both anabolic and catabolic metabolism.

The production of ATP is one of the most important reactions in metabolism.
The synthesis of most ATP is coupled to membrane-associated electron transport
(Chapter 14). In electron transport, the energy of reduced coenzymes such as NADH is
used to generate an electrochemical gradient of protons across a cell membrane. The po-
tential energy of this gradient is harnessed to drive the phosphorylation of ADP to ATP.

ADP + Pj > ATP + H 2 0 (10.4)

We will see that the reactions of membrane- associated electron transport and coupled
ATP synthesis are similar in many ways to the reactions that capture light energy during
photosynthesis (Chapter 15).

Three additional chapters examine the anabolism and catabolism of lipids, amino
acids, and nucleotides. Chapter 16 discusses the storage of nutrient material as triacyl-
glycerols and the subsequent oxidation of fatty acids. This chapter also describes the
synthesis of phospholipids and isoprenoid compounds. Amino acid metabolism is dis-
cussed in Chapter 17. Although amino acids were introduced as the building blocks of
proteins, some also play important roles as metabolic fuels and biosynthetic precursors.
Nucleotide biosynthesis and degradation are considered in Chapter 18. Unlike the other
three classes of biomolecules, nucleotides are catabolized primarily for excretion rather
than for energy production. The incorporation of nucleotides into nucleic acids and of
amino acids into proteins are major anabolic pathways. Chapters 20 to 22 describe these
biosynthetic reactions.

10.4 Compartmentation and Interorgan
Metabolism

Some metabolic pathways are localized to particular regions within a cell. For example,
the pathway of membrane-associated electron transport coupled to ATP synthesis takes
place within the membrane. In bacteria this pathway is located in the plasma membrane
and in eukaryotes it is found in the mitochondrial membrane. Photosynthesis is
another example of a membrane-associated pathway in bacteria and eukaryotes.

10.4 Compartmentation and Interorgan Metabolism 305

Golgi apparatus P
(end-on view)
sorting and secretion
of some proteins

Lysosome:

degradation of proteins
lipids, etc.

Plasma membrane

Mitochondria:

citric acid cycle,
electron transport +
ATP synthesis, fatty
acid degradation

Cytosol:

fatty acid synthesis,
glycolysis, most
gluconeogme:s reaction
pentose phosphase
pathwwary

Nucleus:

nucleic acid synthesis

Endoplasmic reticulum:
delivery of proteins and
synthesis of lipids for
membranes

Nuclear membranes

Figure 10.10 ▲

Compartmentation of metabolic processes within a eukaryotic cell. This is a colored electron micro-
graph of a cell showing the nucleus (green), mitochondria (purple), lysosomes (brown), and exten-
sive endoplasmic reticulum (blue). (Not all pathways and organelles are shown.)

In eukaryotes, metabolic pathways are localized within several membrane-bound
compartments (Figure 10.10). For example, the enzymes that catalyze fatty acid synthe-
sis are located in the cytosol, whereas the enzymes that catalyze fatty acid breakdown
are located inside mitochondria. One consequence of compartmentation is that sepa-
rate pools of metabolites can be found within a cell. This arrangement permits the
simultaneous operation of opposing metabolic pathways. Compartmentation can also
offer the advantage of high local concentrations of metabolites and coordinated regula-
tion of enzymes. Some of the enzymes that catalyze reactions in mitochondria (which
have evolved from a symbiotic prokaryote) are encoded by mitochondrial genes; this
origin explains their compartmentation.

There is also compartmentation at the molecular level. Enzymes that catalyze some
pathways are physically organized into multienzyme complexes (Section 5.11). With
these complexes, channeling of metabolites prevents their dilution by diffusion. Some
enzymes catalyzing adjacent reactions in pathways are bound to membranes and can
diffuse rapidly in the membrane for interaction.

Individual cells of multicellular organisms maintain different concentrations of
metabolites, depending in part on the presence of specific transporters that facilitate the
entry and exit of metabolites. In addition, depending on the cell-surface receptors and
signal-transduction mechanisms present, individual cells respond differently to external
signals.

In multicellular organisms, compartmentation can also take the form of specializa-
tion of tissues. The division of labor among tissues allows site-specific regulation of
metabolic processes. Cells from different tissues are distinguished by their complement
of enzymes. We are very familiar with the specialized role of muscle tissue, red blood
cells, and brain cells but cell compartmentation is a common feature even in simple
species. In cyanobacteria, for example, the pathway for nitrogen fixation is sequestered
in special cells called heterocysts (Figure 10.11). This separation is necessary because
nitrogenase is inactivated by oxygen and the cells that carry out photosynthesis produce
lots of oxygen.

▲ Figure 10.1 1

Anabaena spherica. Many species of cyanobac-
teria form long, multicellular filaments. Some
specialized cells have adapted to carry out
nitrogen fixation. These heterocysts have
become rounded and are surrounded by a
thickened cell wall. The heterocysts are con-
nected to adjacent cells by internal pores.
The formation of heterocysts is an example
of compartmentation of metabolic pathways.

306

CHAPTER 10 Introduction to Metabolism

10.5 Actual Gibbs Free Energy Change, Not
Standard Free Energy Change, Determines
the Direction of Metabolic Reactions

The Gibbs free energy change is a measure of the energy available from a reaction (Sec-
tion 1.4B). The standard Gibbs free energy change for any given reaction (AG°' react i on ) is
the change under standard conditions of pressure (1 atm), temperature (25°C = 298 K),
and hydrogen ion concentration (pH = 7.0). The concentration of every reactant and
product is 1 M under standard conditions. For biochemical reactions, the concentration
of water is assumed to be 55 M.

The standard Gibbs free energy change in a reaction can be determined by using
tables that list the Gibbs free energies of formation (AfG°') of important biochemical
molecules.

AG° reaction AfG° products AfG° reactants ( 10 . 5 )

Keep in mind that Equation 10.5 only applies to the free energy change under standard
conditions where the concentrations of products and reactants are 1 M. It’s also impor-
tant to use tables that apply to biochemical reactions. These tables correct for pH and
ionic strength. The Gibbs free energies of formation under cellular conditions are often
quite different from the ones used in chemistry and physics.

The actual Gibbs free energy change (AG) for a reaction depends on the real con-
centrations of reactants and products, as described in Section 1.4B. The relationship
between the standard free energy change and the actual free energy change is given by

reaction

AG°'

reaction

+ snn lproductsl

[reactants]

( 10 . 6 )

For a chemical or physical process, the free energy change is expressed in terms of the
changes in enthalpy (heat content) and entropy (randomness) as the reactants are con-
verted to products at constant pressure and volume.

AG = AH - TAS

( 10 . 7 )

AH is the change in enthalpy, AS is the change in entropy, and T is the temperature in
degrees Kelvin.

When AG for a reaction is negative, the reaction will proceed in the direction it is
written. When AG is positive, the reaction will proceed in the reverse direction — there will
be a net conversion of products to reactants. For such a reaction to proceed in the direc-
tion written, enough energy must be supplied from outside the system to make the free
energy change negative. When AG is zero, the reaction is at equilibrium and there is no
net synthesis of product.

Because changes in both enthalpy and entropy contribute to AG, the sum of these
contributions at a given temperature (as indicated in Equation 10.7) must be negative for
a reaction to proceed. Thus, even if AS for a particular process is negative (i.e., the prod-
ucts are more ordered than the reactants), a sufficiently negative AH can overcome the
decrease in entropy, resulting in a AG that is less than zero. Similarly, even if AH is posi-
tive (i.e., the products have a higher heat content than the reactants), a sufficiently
positive AS can overcome the increase in enthalpy, resulting in a negative AG. Reactions
that proceed because of a large positive AS are said to be entropy driven. Examples of
entropy- driven processes include protein folding (Section 4.10) and the formation
of lipid bilayers (Section 9.8A), both of which depend on the hydrophobic effect
(Section 2.5D). The processes of protein folding and lipid-bilayer formation result in
states of decreased entropy for the protein molecule and bilayer components, respec-
tively. However, the decrease in entropy is offset by a large increase in the entropy of
surrounding water molecules.

For any enzymatic reaction within a living organism, the actual free energy change
(the free energy change under cellular conditions) must be less than zero in order for

10.5 Actual Gibbs Free Energy Change, Not Standard Free Energy Change, Determines the Direction of Metabolic Reactions 307

the reaction to occur in the direction it is written. Many metabolic reactions have
standard Gibbs free energy changes (AG°' react i on ) that are positive. The difference
between AG and A G°' depends on cellular conditions. The most important condition
affecting free energy change in cells is the concentrations of substrates and products of a
reaction. Consider the reaction

A + B C + D

( 10 . 8 )

At equilibrium, the ratio of substrates and products is by definition the equilibrium
constant (K e q ) and the Gibbs free energy change under these conditions is zero.

[C][D]

(at equilibrium) K e q = AG = 0 (10.9)

When this reaction is not at equilibrium, a different ratio of products to substrates is
observed and the Gibbs free energy change is derived using Equation 10.6.

[C][D]

A (^reaction — AG° reaction RT\f\ faitdi — AG° reaction + RT \f I Q

[A][B]

where q =

[A][B] J

( 10 . 10 )

Q is the mass action ratio. The difference between this ratio and the ratio of products to
substrates at equilibrium determines the actual Gibbs free energy change for a reaction.
In other words, the free energy change is a measure of how far from equilibrium the
reacting system is operating. Consequently, AG, not A G°\ is the criterion for assessing
the direction of a reaction in a biological system.

We can divide metabolic reactions into two types. Let Q represent the steady-state
ratio of product and reactant concentrations in a living cell. Reactions for which Q is
close to K eq are called near-equilibrium reactions. The free energy changes associated with
near- equilibrium reactions are small, so these reactions are readily reversible. Reactions
for which Q is far from K eq are called metabolically irreversible reactions. These reactions
are greatly displaced from equilibrium, with Q usually differing from K eq by two or
more orders of magnitude. Thus, AG is a large negative number for metabolically irre-
versible reactions.

When flux through a pathway changes by a large amount, there may be short-term
perturbations of metabolite concentrations in the pathway. The intracellular concentra-
tions of metabolites vary, but usually over a range of not more than two- or threefold
and equilibrium is quickly restored. As mentioned above, this is called the steady state
condition and it’s typical of most of the reactions in a pathway. Most enzymes in a path-
way catalyze near- equilibrium reactions and have sufficient activity to quickly restore
concentrations of substrates and products to near-equilibrium conditions. They can ac-
commodate flux in either direction. The Gibbs free energy change for these reactions is
effectively zero.

In contrast, the activities of enzymes that catalyze metabolically irreversible reac-
tions are usually insufficient to achieve near-equilibrium status for the reactions. Meta-
bolically irreversible reactions are generally the control points of pathways, and the
enzymes that catalyze these reactions are usually regulated in some way. In fact, the reg-
ulation maintains metabolic irreversibility by preventing the reaction from reaching
equilibrium. Metabolically irreversible reactions can act as bottlenecks in metabolic
traffic, helping control the flux through reactions further along the pathway.

Near-equilibrium reactions are not usually suitable control points. Flux through a
near- equilibrium step cannot be significantly increased since it is already operating
under conditions where the concentrations of products and reactants are close to the
equilibrium values. The direction of near-equilibrium reactions can be controlled by
changes in substrate and product concentrations. In contrast, flux through metabolically
irreversible reactions is relatively unaffected by changes in metabolite concentration; flux
through these reactions must be controlled by modulating the activity of the enzyme.

KEY CONCEPT

Metabolically irreversible reactions are
catalyzed by enzymes whose activity is
regulated in order to prevent the reaction
from reaching equilibrium.

Consider a sample reaction X = Y
under standard conditions of pressure,
temperature, and concentration.

Assume that A G°'

is negative.

•

-o

1 M

A G°' negative

Inside the cell, the reaction will likely
be at equilibrium and AG- 0

X Y

AG=0

(A G°' negative)

For a reaction in which A G°' is
positive,

X Y

o-o

1 M 1 M

A G°' positive

at equilibrium, the concentration of
reactant will be higher than that of the
product.

X Y

AG=0

(A G°' positive)

The standard Gibbs free energy change
does not predict whether a reaction
will proceed in one direction or another.
Instead, it predicts the steady state
concentrations of reactants and prod-
ucts in near-equilibrium reactions.

308

CHAPTER 10 Introduction to Metabolism

Because so many metabolic reactions are near- equilibrium reactions, we have cho-
sen not to emphasize A G°' values in our discussions of most reactions. Those values are
not relevant except when they are used to calculate steady state concentrations.

SAMPLE CALCULATION 10.1 Calculating Standard Gibbs Free Energy Change

from Energies of Formation

For any reaction, the standard Gibbs free energy change for the reaction is given by

AG reaction = AfG° products AfG° reactants
For the oxidation of glucose,

(CH 2 0) 6 + 60 2 -> 6C0 2 + 6H 2 0

you obtain the standard Gibbs free energies of formation from biochemical tables.

AfG°'(glucose) = -426 kj mol -1
A f C°'(0 2 ) = 0
A f G°'(C0 2 ) = -394 kj mol -1
A f G°'(H 2 0) = -156 kj mol -1
AG°' re action = 6(-394) + 6(-1 56) - (-426)

= -2874 kj mol -1

Glucose is an energy- rich organic molecule and its oxidation releases a great deal
of energy. Nevertheless, all living cells routinely synthesize glucose from simple
precursors. In many cases, the precursors are C0 2 and H 2 0 in the reverse of the
reaction shown here. How do they do it?

Section 7.2 A described the structure
and functions of nucleoside triphos-
phates.

Another example of the role of
pyrophosphate is discussed in Sec-
tion 10.7C. Hydrolysis of pyrophos-
phate is often counted as one ATP
equivalent in terms of energy currency.

Table 10.1 Free Energies of Formation
(A f G°')

k) mol 1

ATP

-2102

ADP

-1231

AMP

-360

-1 059

h 2 o

-156

(1 mM Mg®, ionic strength of 0.25 M)

10.6 The Free Energy of ATP Hydrolysis

ATP contains one phosphate ester formed by linkage of the a- phosphoryl group to the
5 '-oxygen of ribose and two phosphoanhydrides formed by the a,/3 and /3, /linkages be-
tween phosphoryl groups (Figure 10.12). ATP is a donor of several metabolic groups,
usually a phosphoryl group, leaving ADP, or an AMP group, leaving inorganic
pyrophosphate (PPi). Both reactions require the cleavage of a phosphoanhydride link-
age. Although the various groups of ATP are not transferred directly to water, hydrolytic
reactions provide useful estimates of the Gibbs free energy changes involved. Table 10.1
lists the free energies of formation of the various reactants and products under standard
conditions, 1 mM Mg 2+ , and an ionic strength of 0.25 M. Table 10.2 lists the standard
Gibbs free energies of hydrolysis ( A G°' hydrolysis) for ATP and AMP, and Figure 10.9
shows the hydrolytic cleavage of each of the phosphoanhydrides of ATP. Note from
Table 10.2 that cleavage of the ester releases only 13 kj mol -1 under standard conditions
but cleavage of either of the phosphoanhydrides releases at least 30 kj mol -1 under stan-
dard conditions.

Table 10.2 also gives the standard Gibbs free energy change for hydrolysis of
pyrophosphate. All cells contain an enzyme called pyrophosphatase that catalyzes this
reaction. The cellular concentration of pyrophosphate is maintained at a very low con-
centration as a consequence of this highly favorable reaction. This means that the
hydrolysis of ATP to AMP + pyrophosphate will always be associated with a negative
Gibbs free energy change even when the AMP concentration is significant.

Nucleoside diphosphates and triphosphates in both aqueous solution and at the ac-
tive sites of enzymes are usually present as complexes with magnesium (or sometimes
manganese) ions. These cations coordinate with oxygen atoms of the phosphate groups,
forming six-membered rings. A magnesium ion can form several different complexes
with ATP; the complexes involving the a and (3 and the /3 and /phosphate groups are
shown in Figure 10.13. Formation of the (3, y complex is favored in aqueous solutions.
We will see later that nucleic acids are also usually complexed with counterions such as

10.6 The Free Energy of ATP Hydrolysis 309

°o— P — o— p-

,©

Adenosine 5' -triphosphate (ATP®)

0 O — P — O — P — O — Adenosine

,©

i©

0 O — P — O — Adenosine

,©

Adenosine 5'-diphosphate (ADP®) Adenosine 5'-monophosphate (AMP®)

◄ Figure 10.12

Hydrolysis of ATP to (1 ) ADP and inorganic
phosphate (Pj) and (2) AMP and inorganic
pyrophosphate (PPj).

The release of a free proton in these
reactions depends on the conditions
since the pKa values of the various
components are close to the value
inside cells (see Figure 2.19).

11 ©
HO— P — 0°

,©

o o

II II Q

HO— P— O— P — 0°

o 0 0°

Inorganic phosphate (Pj)

Inorganic pyrophosphate (PPj)

Mg® or cationic proteins. For convenience, we usually refer to the nucleoside triphos-
phates as adenosine triphosphate (ATP), guanosine triphosphate (GTP), cytidine
triphosphate (CTP), and uridine triphosphate (UTP), but remember that these mole-
cules actually exist as complexes with Mg® in cells.

Several factors contribute to the large amount of energy released during hydrolysis
of the phosphoanhydride linkages of ATP.

1. Electrostatic repulsion. Electrostatic repulsion among the negatively charged oxy-
gen atoms of the phosphoanhydride groups of ATP is less after hydrolysis. [In cells,
AG°' hydrolysis is actually increased (made more positive) by the presence of Mg®,
which partially neutralizes the charges on the oxygen atoms of ATP and diminishes
electrostatic repulsion.]

2. Solvation effects. The products of hydrolysis, ADP and inorganic phosphate, or
AMP and inorganic pyrophosphate, are better solvated than ATP itself. When ions

Table 10.2 Standard Gibbs free energies
of hydrolysis for ATP, AMP, and
pyrophosphate

Reactants
and products

AG 0 ' h y drO |y S i S

(kj mol 1 )

ATP + H 2 0
ADP + Pj + H®

-32

ATP + H 2 0 ->
AMP + PP; + H®

-45

AMP + H 2 0^
Adenosine + P; + H®

-13

PPj + H 2 0 — » 2Pj

-29

Pj(inorganic phosphate) =

hpo 4 ©

PPj(pyrophosphate) = HP 2 0 7 ®

0 0 0

~ II II II

^O — P y — O — Pp — O — P a — O — Adenosine a, (3 complex of MgATP

©o ®o, .0©

0 0 0

O II II II

^O — P„ — O — P B — O — P a — O — Adenosine

f i i

©o. ,o© o©

/ 3 , y complex of MgATP

◄ Figure 10.13

Complexes between ATP and Mg©.

310 CHAPTER 10 Introduction to Metabolism

A quantitative definition of a “high
energy” compound is presented in
Section 10.7A.

KEY CONCEPT

The large free energy change associated
with hydrolysis of ATP is only possible if
the system is far from equilibrium.

Table 10.3 Theoretical changes in

concentrations of adenine
nucleotides

ATP

ADP

AMP

(mM)

4.8

0.2

0.004

4.5

0.5

0.02

3.9

1.0

0.11

3.2

1.5

0.31

[Adapted from Newsholme. E. A., and Leech, A. R.
(1 986). Biochemistry for the Medical Science (New
York: John Wiley & Sons), p. 315.]

are solvated, they are electrically shielded from each other. Solvation effects are
probably the most important factor contributing to the energy of hydrolysis.

3. Resonance stabilization. The products of hydrolysis are more stable than ATP.
The electrons on terminal oxygen atoms are more delocalized than those on bridg-
ing oxygen atoms. Hydrolysis of ATP replaces one bridging oxygen atom with two
new terminal oxygen atoms.

Because of the free energy change associated with the cleavage of their phosphoan-
hydrides, ATP and the other nucleoside triphosphates (UTP, GTP, and CTP) are often
referred to as energy-rich compounds, but keep in mind that its the system, not the mole-
cule, that contributes free energy to biochemical reactions. ATP, by itself, is not really a
high energy compound. It can only work if the system (reactants and products) is far
from equilibrium. The ATP currency becomes worthless if the reaction reaches equilib-
rium and AG = 0. We will find it useful to refer to “energy-rich” or “high energy” mole-
cules in the jargon of biochemistry but we will put the terms in quotation marks to re-
mind you that it is jargon.

All the phosphoanhydrides of nucleoside triphosphates have nearly equal standard
Gibbs free energies of hydrolysis. We occasionally express the consumption or formation
of the phosphoanhydride linkages of nucleoside triphosphates in terms of ATP equivalents.

ATP is usually the phosphoryl group donor when nucleoside monophosphates and
diphosphates are phosphorylated. Of course, the intracellular concentrations of indi-
vidual nucleoside mono-, di-, and triphosphates differ, depending on metabolic needs.
For example, the intracellular levels of ATP are far greater than deoxythymidine
triphosphate (dTTP) levels. ATP is involved in many reactions, whereas dTTP has fewer
functions and is primarily a substrate for DNA synthesis.

A series of kinases (phosphotransferases) catalyze interconversions of nucleoside
mono-, di-, and triphosphates. Phosphoryl group transfers between nucleoside phos-
phates have equilibrium constants close to 1.0. Nucleoside monophosphate kinases are
a group of enzymes that catalyze the conversion of nucleoside monophosphates to
nucleoside diphosphates. For example, guanosine monophosphate (GMP) is converted
to guanosine diphosphate (GDP) by the action of guanylate kinase. GMP or its deoxy
analog dGMP is the phosphoryl group acceptor in the reaction, and ATP or dATP is the
phosphoryl group donor.

GMP + ATP GDP + ADP (10.11)

Nucleoside diphosphate kinase acts in the conversion of nucleoside diphosphates
to nucleoside triphosphates. This enzyme, present in both the cytosol and mitochondria
of eukaryotes, is much less specific than nucleoside monophosphate kinases. All nucleo-
side diphosphates, regardless of the purine or pyrimidine base, are substrates for nucle-
oside diphosphate kinase. Nucleoside monophosphates are not substrates. Because of
its relative abundance, ATP is usually the phosphoryl- group donor in cells:

GDP + ATP GTP + ADP (10.12)

Although the concentration of ATP varies among cell types, the intracellular ATP
concentration fluctuates very little within a particular cell, and the sum of the concen-
trations of the adenine nucleotides remains nearly constant. Intracellular ATP concen-
trations are maintained in part by the action of adenylate kinase that catalyzes the fol-
lowing near- equilibrium reaction:

AMP + ATP 2 ADP (10.13)

When the concentration of AMP increases, AMP can react with ATP to form two mole-
cules of ADP. These ADP molecules can be converted to two molecules of ATP. The
overall process is

AMP + ATP + 2 Pi 2 ATP + 2 H 2 0 (10.14)

ATP concentrations in cells are greater than ADP or AMP concentrations, and rela-
tively minor changes in the concentration of ATP can result in large changes in the con-
centrations of the di- and monophosphates. Table 10.3 shows the theoretical increases in

10.6 The Free Energy of ATP Hydrolysis

311

[ADP] and [AMP] under conditions in which ATP is consumed, assuming that the total
adenine nucleotide concentration remains 5.0 mM. Note that when the ATP concentra-
tion decreases from 4.8 mM to 4.5 mM (a decrease of about 6%), the ADP concentration
increases 2.5-fold and the AMP concentration increases 5-fold. In fact, when cells are well
supplied with oxidizable fuels and oxygen, they maintain a balance of adenine nucleotides
in which ATP is present at a steady concentration of 2 to 10 mM, [ADP] is less than 1 mM,
and [AMP] is even lower. As we will see, ADP and AMP are often effective allosteric mod-
ulators of some energy-yielding metabolic processes. ATP, whose concentration is rela-
tively constant, is generally not an important modulator under physiological conditions.

One important consequence of the concentrations of ATP and its hydrolysis prod-
ucts in vivo is that the free energy change for ATP hydrolysis is actually greater than the
standard value of —32 kj mol -1 . This is illustrated in Sample Calculation 10.2 using
measured concentrations of ATP, ADP, and Pi from rat liver cells. The calculated Gibbs
free energy change is close to the value determined in many other types of cells.

As mentioned above, ATP hydrolysis is an example of a metabolically irreversible
reaction. The activities of various enzymes are regulated so they become inactive as ATP
concentrations fall below a minimal threshold. Thus, the reverse of the hydrolysis reac-
tion, leading to ATP synthesis, does not occur except under special circumstances
(Chapter 14). We will see in Chapter 14 that ATP is synthesized by another pathway.

The importance of maintaining a high concentraion of ATP cannot be overempha-
sized. It is required in order to get a large free energy change from ATP hydrolysis. Cells
will die if the reactants and products reach equilibrium.

10.7 The Metabolic Roles of ATP

The energy produced by one biological reaction or process, such as the synthesis of
X — Y in Reaction 10.15, is often coupled to a second reaction, such as the hydrolysis
of ATP. The first reaction would not otherwise occur spontaneously.

X + Y X— Y

ATP + H 2 0 ADP + Pi + H© (10.15)

SAMPLE CALCULATION 10.2 Gibbs Free Energy Change

Q: In a rat hepatocyte, the concentrations of ATP, ADP, and the Gibbs free energy change for hydrolysis of ATP in this cell.

Pi are 3.4 mM, 1.3 mM, and 4.8 mM, respectively. Calculate How does this compare to the standard free energy change?

A: The actual Gibbs free energy change is calculated according to Equation 10.10.

[ADP][Pj] [ADP][Pj]

Abreaction - AG° reac tj on + /?7ln — AG° reac tj on + 2.303 RTlog

[ATP] [ATP]

When known values and constants are substituted (with concentrations expressed as molar values), assuming pH7.0 and 25°C.

. i i r (1.3 x 10 _3 )(4.8 x 10 -3 )

AC = -32000 j moP 1 + (8.31 JK“ 1 mor 1 )(298 K) 2.303 log r

L (3.4 X 10“ 3 )

AC = -32000 j mol -1 + (2480 j mol" 1 ) [2.303 log(1.8 X 10“ 3 )]

AC = -32000 j moP 1 - 16000 j mol -1
AG = -48000 j mol -1 = -48 kj mol -1

The actual free energy change is about lV 2 times the standard free energy change.

312

CHAPTER 10 Introduction to Metabolism

The sum of the Gibbs free energy changes for the coupled reactions must be negative
for the reactions to proceed. This does not mean that both of the individual reactions
have to be favored in isolation (AG < 0). The advantage of coupled reactions is that the
energy released from one of them can be used to drive the other even when the second
reaction is unfavorable by itself (AG > 0). (Recall that the ability to couple reactions is
one of the key properties of enzymes.)

Energy flow in metabolism depends on many coupled reactions involving ATR In
many cases, the coupled reactions are linked by a shared intermediate such as a phos-
phorylated derivative of reactant X.

X + ATP X — P + ADP

x — P+Y+H 2 0 ^^ x— Y + Pi + H© (10.16)

Transfer of either a phosphoryl group or a nucleotidyl group to a substrate
activates that substrate (i.e., prepares it for a reaction that has a large negative Gibbs
free energy change). The activated compound (X — P), can be either a metabolite or
the side chain of an amino acid residue in the active site of an enzyme. The intermediate
then reacts with a second substrate to complete the reaction.

A. Phosphoryl Group Transfer

The synthesis of glutamine from glutamate and ammonia illustrates how the “high
energy” compound ATP drives a biosynthetic reaction. This reaction, catalyzed by glut-
amine synthetase, allows organisms to incorporate inorganic nitrogen into biomolecules
as carbon-bound nitrogen. In this synthesis of an amide bond, the y-carboxyl group of
the substrate is activated by synthesis of an anhydride intermediate.

Glutamine synthetase catalyzes the nucleophilic displacement of the y-phosphoryl
group of ATP by the y-carboxylate of glutamate. ADP is released, producing enzyme-
bound y-glutamyl phosphate as an intermediate (Figure 10.14). y-Glutamyl phosphate is
unstable in aqueous solution but is protected from water in the active site of glutamine
synthetase. In the second step of the mechanism, ammonia acts as a nucleophile, dis-
placing the phosphate (a good leaving group) from the carbonyl carbon of y-glutamyl
phosphate to generate the product, glutamine. Overall, one molecule of ATP is converted
to ADP + Pj for every molecule of glutamine formed from glutamate and ammonia.

BOX 10.1 THESQUIGGLE

Fritz Fipmann (1899-1986) won the Nobel Prize in Physiology and Medicine in 1953
for discovering coenzyme A. He also made important contributions to our under-
standing of ATP as an energy currency. In 1941 he introduced the idea of a high
energy bond in ATP by drawing it as a squiggle (~). For the next several decades,
biochemistry textbooks often depicted ATP with two high energy bonds.

AMP~P~P

We know now that this depiction is misleading since there’s nothing special
about the covalent bonds in phosphoanhydride linkages. It’s the overall system of re-
actants and products that makes the ATP currency so valuable and not the energy of
individual bonds. However, it’s true that the three main explanations for the high en-
ergy of ATP (electrostatic repulsion, solvation effects, and resonance stabilization) are
due mostly to the phosphoanhydride linkages so the focus on that particular linkage
isn’t entirely wrong. The squiggle used to be very common in the older scientific liter-
ature and in textbooks but it’s much less common today.

Source: Lipmann, F. (1941) Metabolic generation and utilization of phosphate bond energy. Advances in
Enzymology 1:99-162.

10.7 The Metabolic Roles of ATP

313

COO

:NI-U

HoN— C— H

CH 2

ch 2

f \

o nh 2

Glutamine

( 10 . 17 )

We can calculate the predicted standard Gibbs free energy change for the reaction
that is not coupled to ATP hydrolysis.

Glutamate + NH^ glutamine + H 2 0 (10.18)

AG reaction — +14 kj mol

This is a standard free energy change so it doesn’t necessarily reflect the actual Gibbs
free energy change given cellular concentrations of glutamate, glutamine, and ammo-
nia. The hypothetical Reaction 10.18 might be associated with a negative free energy
change inside the cell if the concentrations of glutamate and ammonia were high rela-
tive to the concentration of glutamine. But this is not the case. The steady- state concen-
trations of glutamate and glutamine must be kept nearly equivalent in order to support
protein synthesis and other metabolic pathways. This means that the Gibbs free energy
change for the hypothetical Reaction 10.18 cannot be negative. Furthermore, the con-
centration of ammonia is very low relative to glutamate and glutamine. In both bacteria
and eukaryotes, ammonia must be efficiently incorporated into glutamine even when
the concentration of free ammonia is very low. Thus Reaction 10.18 is not possible in
living cells due to the requirement for a high steady- state concentration of glutamine
and due to a limiting supply of ammonia. Glutamine synthesis must be coupled to
hydrolysis of ATP in order to drive it in the right direction.

Glutamine synthetase catalyzes a phosphoryl group transfer reaction in which the
phosphorylated compound is a transient intermediate (Reaction 10.17). There are other
reactions that produce a stable phosphorylated product. As we have seen, kinases catalyze

◄ Figure 10.14

Glutamine synthetase bound to ADP and a tran-
sition state analog. Glutamine synthetase
from Mycobacterium tuberculosis is a com-
plex enzyme consisting of two hexameric
rings on top of each other. Only one ring is
shown in this figure. The active site is occu-
pied by ADP and a transition state analog
(L-methionine-S-sulfoximine phosphate) that
resembles y-glutamyl phosphate.

[PDB 2BVC]

314 CHAPTER 10 Introduction to Metabolism

Table 10.4 Standard Gibbs free energies
of hydrolysis for common
metabolites

Metabolite

A C hydrolysis

(kj mol -1 )

Phosphoenolpyruvate

-62

1, 3-8/sphosphoglycerate

-49

ATP to AMP + PPj

-45

Phosphocreatine

-43

Phosphoarginine

-32

Acetyl CoA

-32

Acyl CoA

-31

ATP to ADP + Pi

-32

Pyrophosphate

-29

Glucose 1 -phosphate

-21

Glucose 6-phosphate

-14

Glycerol 3-phosphate

-9

KEY CONCEPT

Many phosphorylated metabolites have
group transfer potentials similar to that
of ATP.

transfer of the y-phosphoryl group from ATP (or, less frequently, from another nucleo-
side triphosphate) to another substrate. Kinases typically catalyze metabolically irre-
versible reactions. A few kinase reactions, however, such as those catalyzed by adenylate
kinase (Reaction 10.13) and creatine kinase (Section 10.7B), are near equilibrium reactions.
Although the reactions they catalyze are sometimes described as phosphate group trans-
fer reactions, kinases actually transfer a phosphoryl group ( — P0 3 ^ — )to their acceptors.

The ability of a phosphorylated compound to transfer its phosphoryl group (s) is
termed its phosphoryl group transfer potential, or simply group transfer potential. Some
compounds, such as phosphoanhydrides, are excellent phosphoryl group donors. They
may have a group transfer potential equal to or greater than that of ATP. Other com-
pounds, such as phosphoesters, are poor phosphoryl group donors. They have a group
transfer potential less than that of ATP. Under standard conditions, group transfer poten-
tials have the same values as the standard free energies of hydrolysis but are opposite in
sign. Thus, the group transfer potential is a measure of the free energy required for for-
mation of the phosphorylated compound. In Table 10.4 we list the standard Gibbs free
energy of hydrolysis for a number of phosphorylated compounds.

B. Production of ATP by Phosphoryl Group Transfer

Often, one kinase catalyzes transfer of a phosphoryl group from an excellent donor to
ADP to form ATP, which then acts as a donor for a different kinase reaction. Phospho-
enolpyruvate and 1,3-bisphosphoglycerate are two examples of common metabolites
that have higher energy than ATP even under conditions found inside the cell (AG <
—50 kj mol -1 ). Some of these compounds are intermediates in catabolic pathways; oth-
ers are energy storage compounds.

Phosphoenolpyruvate, an intermediate in the glycolytic pathway, has the highest
phosphoryl group transfer potential known. The standard free energy of phospho-
enolpyruvate hydrolysis is —62 kj mol -1 and the actual Gibbs free energy change is com-
parable to that of ATP. The free energy of hydrolysis for phosphoenolpyruvate can be
understood by picturing the molecule as an enol whose structure is locked by attach-
ment of the phosphoryl group. When the phosphoryl group is removed, the molecule
spontaneously forms the much more stable keto tautomer (Figure 10.15). Transfer of
the phosphoryl group from phosphoenolpyruvate to ADP is catalyzed by the enzyme
pyruvate kinase. Because the A G°' for the reaction is about —30 kj mol -1 , the equilib-
rium for this reaction under standard conditions lies far in the direction of transfer of
the phosphoryl group from phosphoenolpyruvate to ADP. In cells, this metabolically
irreversible reaction is an important source of ATP.

Phosphagens, including phosphocreatine and phosphoarginine, are “high energy”
phosphate storage molecules found in animal muscle cells. Phosphagens are phospho-
amides (rather than phosphoanhydrides) and have higher group transfer potentials
than ATP. In the muscles of vertebrates, large amounts of phosphocreatine are formed
during times of ample ATP supply. In resting muscle, the concentration of phosphocre-
atine is about fivefold higher than that of ATP. When ATP levels fall, creatine kinase cat-
alyzes rapid replenishment of ATP through transfer of the activated phosphoryl group
from phosphocreatine to ADP.

Creatine

kinase

Phosphocreatine + ADP creatine + ATP (10.19)

The supply of phosphocreatine is adequate for 3- to 4-second bursts of activity, long enough
for other metabolic processes to begin restoring the ATP supply. Under cellular conditions,
the creatine kinase reaction is a near-equilibrium reaction. In many invertebrates —
notably mollusks and arthropods — phosphoarginine is the source of the activated
phosphoryl group.

Because ATP has an intermediate phosphoryl group transfer potential, it is thermo-
dynamically suited as a carrier of phosphoryl groups. (Figure 10.15) ATP is also
kinetically stable under physiological conditions until acted on by an enzyme so it can
carry chemical potential energy from one enzyme to another without being hydrolyzed.
Not surprisingly, ATP mediates most chemical energy transfers in all organisms.

10.7 The Metabolic Roles of ATP 315

COO° 0

ADP ATP

coo 0

1 II 0

c — 0 — P — 0°

w ,

C — OH

II l n

C 0°

Pyruvate kinase

/ \

H H

Phosphoenolpyruvate

Enolpyruvate

COOO ◄Figure 10.15

Transfer of the phosphoryl group from phospho
C = O enolpyruvate to ADP.

H — C — H

Pyruvate

C. Nucleotidyl Group Transfer

The other common group transfer reaction involving ATP is transfer of the nucleotidyl
group. An example is the synthesis of acetyl Co A, catalyzed by acetyl- Co A synthetase. In
this reaction, the AMP moiety of ATP is transferred to the nucleophilic carboxylate
group of acetate to form an acetyl-adenylate intermediate (Figure 10.16). Note that
pyrophosphate (PPi) is released in this step. Like the glutamyl-phosphate intermediate
in Reaction 10.17, the reactive intermediate is shielded from nonenzymatic hydrolysis
by tight binding within the active site of the enzyme. The reaction is completed by
transfer of the acetyl group to the nucleophilic sulfur atom of coenzyme A, leading to
the formation of acetyl CoA and AMP.

The synthesis of acetyl CoA also illustrates how the removal of a product can cause
a metabolic reaction to approach completion, just as the formation of a precipitate or
a gas can drive an inorganic reaction toward completion. The standard Gibbs free
energy for the formation of acetyl CoA from acetate and CoA is about — 13 kj mol -1
(AG°' h y( j ro i 7S i s of acetyl CoA = —32 kj mol -1 ). But note that the product PPj is hy-
drolyzed to two molecules of by the action of pyrophosphatase (Section 10.6). Al-
most all cells have high levels of activity of this enzyme, so the concentration of PPi in
cells is generally very low (less than 10 -6 M). Cleavage of PPi contributes to the negative
value of the standard Gibbs free energy change for the overall reaction. The additional
hydrolytic reaction adds the energy cost of one phosphoanhydride linkage to the overall
synthetic process. In reactions such as this, we say that the cost is two ATP equivalents in
order to emphasize that two “high energy” compounds are hydrolyzed. Hydrolysis of
pyrophosphate accompanies many synthetic reactions in metabolism.

COO

C h 2

HoC — N

©^ \ II

h 2 n n — p— o
H I
o 0

Phosphocreatine

,©

COO'

,©

HoN — C — H

(CH 2 ) 3

©^ \

h 2 n n — p — o'

,©

I©

Phosphoarginine

▲ Structures of phosphocreatine and
phosphoarginine.

Acetate O

11 O
HoC — C— O®

©,

O — P — O — P — O — P — O — Adenosine

(1)

»©

6 °

ATP

»©

c ll II

HoC — C — O — P — O — Adenosine

\©

H 2 0

+ PP,

Pyrophosphatase

* 2P:

H — S-CoA

Enzyme-bound

acetyl-adenylate

intermediate

( 2 )

H ©

H 3 C — C — S-CoA

Acetyl CoA

(3)

AMP

(°0 o

f\ II

H 3 C — C — O — P — O — Adenosine

1 >©

S-CoA O u

▲ Figure 10.16

Synthesis of acetyl CoA from acetate, catalyzed by acetyl-CoA synthetase.

316

CHAPTER 10 Introduction to Metabolism

10.8 Thioesters Have High Free
Energies of Hydrolysis

Thioesters are another class of “high energy” compounds forming part of the currency
of metabolism. Acetyl CoA is one example. It occupies a central position in metabolism
(Figures 10.8 and 10.9). The high energy of thioester reactions can be used in generat-
ing ATP equivalents or in transferring the acyl groups to acceptor molecules. Recall that
acyl groups are attached to coenzyme A (or acyl carrier protein) via a thioester linkage
(Section 7.6 and Figure 7.13).

R — C — S — Coenzyme A

( 10 . 20 )

Unlike oxygen esters of carboxylic acids, thioesters resemble carboxylic acid anhy-
drides in reactivity. Sulfur is in the same group of the periodic table as oxygen but
thioesters are less stable than typical esters because the unshared electrons of the sulfur
atom are not as effectively delocalized in a thioester as the unshared electrons in an oxy-
gen ester. The energy associated with hydrolyzing the thioester linkage is similar to the
energy of hydrolysis of the phosphoanhydride linkages in ATP. The standard Gibbs free
energy change for hydrolysis of acetyl CoA is —31 kj mol -1 , and the actual change may
somewhat smaller (more negative) under conditions inside the cell.

KEY CONCEPT

Reactions involving thioesters, such as
acetyl CoA, release amounts of energy
comparable to that of ATP hydrolysis.

o H 2 0 HS-CoA o

H 3 c — C — S-CoA — ^ T > H 3 C — C — O 0 + H® (10.21)

Acetyl CoA Acetate

Despite its high free energy of hydrolysis, a CoA thioester resists nonenzymatic hydrolysis
at neutral pH values. In other words, it is kinetically stable in the absence of appropriate
catalysts.

The high energy of hydrolysis of a CoA thioester is used in the fifth step of the cit-
ric acid cycle, when the thioester succinyl CoA reacts with GDP (or sometimes ADP)
and Pi to form GTP (or ATP).

coo°

ch 2

+ GDP + Pi <

> 1 +

cn.

oh 2

We discuss succinyl CoA synthetase in

c=o

coo 0

Section 13 . 4 , part 5 , and fatty acid
synthesis in Section 16 . 5 .

S-CoA

Succinate

+ HS-CoA

( 10 . 22 )

Succinyl CoA

This substrate-level phosphorylation conserves energy used in the formation of
succinyl CoA as ATP equivalents. The energy of thioesters also drives the synthesis of
fatty acids.

In Section 14.1 1 we will learn that
NADH is equivalent to 2.5 ATPs and
QH 2 is equivalent to 1.5 ATPs.

10.9 Reduced Coenzymes Conserve Energy
from Biological Oxidations

Many reduced coenzymes are “high energy” compounds in the sense we described ear-
lier (i.e., part of a system). Their high energy (or reducing power) can be donated in
oxidation- reduction reactions. The energy of reduced coenzymes maybe represented as
ATP equivalents since their oxidation can be coupled to the synthesis of ATP.

10.9 Reduced Coenzymes Conserve Energy from Biological Oxidations 317

As described in Section 6. 1C, the oxidation of one molecule must be coupled with
the reduction of another molecule. A molecule that accepts electrons and is reduced is
an oxidizing agent. A molecule that loses electrons and is oxidized is a reducing agent.
The net oxidation-reduction reaction is

Ared + B ox A ox + B rec | (10.23)

The electrons released in biological oxidation reactions are transferred enzymati-
cally to oxidizing agents, usually a pyridine nucleotide (NAD® or sometimes NADP®),
a flavin coenzyme (FMN or FAD), or ubiquinone (Q). When NAD® and NADP® are
reduced, their nicotinamide rings accept a hydride ion (Figure 7.8). One electron is lost
when a hydrogen atom (composed of one proton and one electron) is removed and two
electrons are lost when a hydride ion (composed of one proton and two electrons) is
removed. (Remember that oxidation is loss of electrons.)

NADH and NADPH, along with QH 2 , supply reducing power. FMNH 2 and FADH 2
are reduced enzyme-bound intermediates in some oxidation reactions.

A. Gibbs Free Energy Change Is Related to Reduction Potential

The reduction potential of a reducing agent is a measure of its thermodynamic reactivity.
Reduction potential can be measured in electrochemical cells. An example of a simple
inorganic oxidation-reduction reaction is the transfer of a pair of electrons from a zinc
atom (Zn) to a copper ion (Cu©).

Zn + Cu® Zn® + Cu (10.24)

This reaction can be carried out in two separate solutions that divide the overall reaction
into two half- reactions (Figure 10.17). At the zinc electrode, two electrons are given up
by each zinc atom that reacts (the reducing agent). The electrons flow through a wire to
the copper electrode, where they reduce Cu© (the oxidizing agent) to metallic copper. A
salt bridge, consisting of a tube with a porous partition filled with electrolyte, preserves
electrical neutrality by providing an aqueous path for the flow of nonreactive counteri-
ons between the two solutions. The flow of ions and the flow of electons are separated
in such an electrochemical cell and electron flow through the wire (i.e., electric energy)
can be measured using a voltmeter.

The direction of the current through the circuit in Figure 10.17 indicates that Zn is
more easily oxidized than Cu (i.e., Zn is a stronger reducing agent than Cu). The reading
on the voltmeter represents a potential difference, the difference between the reduction
potential of the reaction on the left and that on the right. The measured potential differ-
ence is the electromotive force.

Voltmeter

The structures and functions of
NAD® and NADP® are discussed
in Section 7.4, of FMN and FAD in
Section 7.5, and of ubiquinone
in Section 7.14.

◄ Figure 10.17

Diagram of an electrochemical cell. Electrons
flow through the external circuit from the
zinc electrode to the copper electrode. The
salt bridge permits the flow of counterions
(sulfate ions in this example) without exten-
sive mixing of the two solutions. The electro-
motive force is measured by the voltmeter
connected across the two electrodes. (Two
other kinds of salt bridges are shown in
Section 2.5A.)

318 CHAPTER 10 Introduction to Metabolism

KEY CONCEPT

All standard reduction potentials are
measured relative to the reduction of H©
under standard conditions.

KEY CONCEPT

A E must be positive for an oxidation
reduction reaction to proceed in the
direction written.

It is useful to have a reference standard for measurements of reduction potentials
just as in measurements of Gibbs free energy changes. For reduction potentials, the ref-
erence is not simply a set of reaction conditions, but a reference half-reaction to which
all other half-reactions can be compared. The reference half- reaction is the reduction of
H© to hydrogen gas (H 2 ). The reduction potential of this half- reaction under standard
conditions (E°) is arbitrarily set at 0.0 V. The standard reduction potential of any other
half- reaction is measured with an oxidation-reduction coupled reaction in which the
reference half-cell contains a solution of 1 M H® and 1 atm H 2 (gaseous), and the sam-
ple half-cell contains 1 M each of the oxidized and reduced species of the substance
whose reduction potential is to be determined. Under standard conditions for biologi-
cal measurements, the hydrogen ion concentration in the sample half-cell is (10 -7 M).
The voltmeter across the oxidation-reduction couple measures the electromotive force,
or the difference in the reduction potential, between the reference and sample half-
reactions. Since the standard reduction potential of the reference half-reaction is 0.0 V,
the measured potential is that of the sample half- reaction.

Table 10.5 gives the standard reduction potentials at pH 7.0 (£°') of some important
biological half- reactions. Electrons flow spontaneously from the more readily oxidized

Table 10.5 Standard reduction potentials of some important biological half-reactions

Reduction half-reaction

£°'(V)

Acetyl CoA + C0 2 + H© + 2e© — » Pyruvate + CoA

0^0

Ferredoxin (spinach). Fe + —> Fe

-0.48

-0.43

2 H© + 2e© -> H 2 (at pH 7.0)

-0.42

a-Ketoglutarate + C0 2 + 2 H© + 2e© — » Isocitrate

-0.38

Lipoyl dehydrogenase (FAD) + 2 H© + 2e© —> Lipoyl dehydrogenase (FADH 2 )

-0.34

NADP© + H© + 2e© — » NADPH

-0.32

NAD© + H© + 2e© — » NADH

-0.32

Lipoic acid + 2 H© + 2e© —> Dihydrolipoic acid

-0.29

Thioredoxin (oxidized) + 2H® + 2e — »Thioredoxin (reduced)

-0.28

Glutathione (oxidized) + 2 H© + 2e©— >2 Glutathione (reduced)

-0.23

FAD + 2 H© + 2e© —> FADH 2

-0.22

FMN + 2 H© + 2e© —> FMNH 2

-0.22

Acetaldehyde + 2 H© + 2e© — » Ethanol

-0.20

Pyruvate + 2 H© + 2e© —> Lactate

-0.18

Oxaloacetate + 2 H© + 2c© —> Malate

0) O r

Cytochrome b 5 (microsomal). Fe + — * Fe

-0.17

0.02

Fumarate + 2 H© + 2e© —> Succinate

0.03

Ubiquinone (Q) + 2 H© + 2e©^QH 2

0.04

0) O r

Cytochrome b (mitochondrial), Fe + - * Fe

0) (0

Cytochrome c-|, Fe + e u — > Fe

0.08

0.22

0 n r- ©

Cytochrome c, Fe + e u —> Fe

(0 (0

Cytochrome a, Fe + > Fe

0.23

0.29

Cytochrome /, Fe + e© —* Fe

0.36

Plastocyanin, Cu 2+ + e©^Cu +

0.37

N0 3 © + 2 H© + 2e© -> N0 2 © + H 2 0

0.42

Photosystem 1 (P700)

0.43

Fe^ + e© — » Fe^

0.77

y 2 0 2 + 2 H© + 2e© — > H 2 0

0.82

Photosystem II (P680)

1.1

10.9 Reduced Coenzymes Conserve Energy from Biological Oxidations

319

substance (the one with the more negative reduction potential) to the more readily
reduced substance (the one with the more positive reduction potential). Therefore,
more negative potentials are assigned to reaction systems that have a greater tendency to
donate electrons (i.e., systems that tend to oxidize more easily).

The standard reduction potential for the transfer of electrons from one molecular
species to another is related to the standard free energy change for the oxidation-reduc-
tion reaction by the equation

A C°' = -nFAE°' (10.25)

where n is the number of electrons transferred and F is Faraday’s constant (96.48 kj
V -1 mol -1 ). Note that Equation 10.25 resembles Equation 9.5 except that here we are
dealing with reduction potential and not membrane potential. A E°' is defined as the
difference in volts between the standard reduction potential of the electron- acceptor
system and that of the electron donor system.

A E ot

= E of

electron acceptor

- E of

electron donor

(10.26)

Recall from Equation 10.6 that A G°' = — RT In K eq . Combining this equation with
Equation 10.25, we have

A E of = —In K eQ (10.27)

nF H

Under biological conditions, the reactants in a system are not present at standard con-
centrations of 1 M. Just as the actual Gibbs free energy change for a reaction is related to
the standard Gibbs free energy change by Equation 10.6, an observed difference in re-
duction potentials (A E) is related to the difference in the standard reduction potentials
(A E°') by the Nernst equation. For Reaction 10.23, the Nernst equation is

A E = A E of

[AoxH^red]

[Aredlt^ox]

(10.28)

At 298 K, Equation 10.28 reduces to

0.026

A£ = A E°’ In Q (10.29)

where Q represents the actual concentrations of reduced and oxidized species. To calcu-
late the electromotive force of a reaction under nonstandard conditions, use the Nernst
equation and substitute the actual concentrations of reactants and products. Keep in
mind that a positive A E value indicates that an oxidation-reduction reaction will have a
negative standard Gibbs free energy change.

B. Electron Transfer from NADH Provides Free Energy

NAD® is reduced to NADH in coupled reactions where electrons are transferred from
a metabolite to NAD®. The reduced form of the coenzyme (NADH) becomes a source
of electrons in other oxidation-reduction reactions. The Gibbs free energy changes as-
sociated with the overall oxidation-reduction reaction under standard conditions can
be calculated from the standard reduction potentials of the two half-reactions using
Equation 10.25. As an example, let’s consider the reaction where NADH is oxidized and
molecular oxygen is reduced. This represents the available free energy change during
membrane-associated electron transport. This free energy is recovered in the form of
ATP synthesis (Chapter 14).

The two half reactions from Table 10.5 are

NAD© + H© + 2 e© * NADH P' = -0.32 V (10.30)

and

y 2 0 2 + 2 H© + 2 e©

* H 2 0 £°' = 0.82 V

(10.31)

320

CHAPTER 10 Introduction to Metabolism

Since the NAD® half- reaction has the more negative standard reduction potential,
NADH is the electron donor and oxygen is the electron acceptor. Note that the values in
Table 10.5 are for half- reactions written as reductions (gain of electrons). That’s because
E°' is a reduction potential. In an oxidation-reduction reaction, two of these half-
reactions are combined. One of them will be an oxidation reaction, so the equation in
Table 10.5 must be reversed. The reduction potentials tell you which way the electrons
will flow. They flow from the half-reaction near the top of the table (more negative E°')
to the one nearer the bottom of the table (less negative E°') (Figure 10.18). What this
means is that the overall A E or for the complete reaction will be positive according to
Equation 10.26. (This is the American convention. The European convention uses a dif-
ferent way of arriving at the same answer.)

The net oxidation-reduction reaction is Reaction 10.31 plus the reverse of
Reaction 10.30.

NADH + % 0 2 + H© > NAD© + H 2 Q (10.32)

and A E°’ for the reaction is

A E of = E% 2 - £ft ADH = 0.82 V - (-0.32 V) = 1.14 V (10.33)

Using Equation 10.25,

AC°' = -(2) (96.48 kj V -1 mol“ 1 )(1.14V) = -220 kj mol -1 (10.34)

KEY CONCEPT

The standard Gibbs free energy change
of an oxidation-reduction reaction is
calculated from the reduction potentials
of the two half-reactions.

The standard Gibbs free energy change for the formation of ATP from ADP + Pj is
+32 kj mol -1 (the actual free energy change is about +48 kj mol -1 under the conditions
of the living cell, as noted earlier). The energy released during the oxidation of NADH
under cellular conditions is sufficient to drive the formation of several molecules of
ATP. We will learn in Chapter 14 that the actual energy yield of an NADH molecule is
about 2.5 ATP equivalents (Section 14.11).

Figure 10.18 ►

Electron flow in oxidation-reduction reactions.

Half-reactions can be plotted on a chart
where the standard reduction potentials are
on the x axis, arranged so that the most neg-
ative values are at the top of the chart.

Using this convention, electrons flow from
the half-reaction at the top of the chart to
the one nearer the bottom of the chart.

10.10 Experimental Methods for Studying Metabolism 321

BOX 10.2 NAD© AND NADH DIFFER IN THEIR ULTRAVIOLET ABSORPTION SPECTRA

The differing absorption spectra of NAD® and NADH are
useful in experimental work. NAD® (and NADP® ) absorbs
maximally at 260 nm. This absorption is due to both the ade-
nine and nicotinamide moieties. When NAD® is reduced to
NADH (or NADP® to NADPH), the absorbance at 260 nm
decreases and an absorption band centered at 340 nm appears
(adjacent figure). The 340-nm band comes from the formation
of the reduced nicotinamide ring. The spectra of NAD® and
NADH do not change in the pH range 2 to 10 in which most
enzymes are active. In addition, few other biological molecules
undergo changes in light absorption near 340 nm.

In a suitably prepared enzyme assay, one can determine
the rate of formation of NADH by measuring an increase in
the absorbance at 340 nm. Similarly, in a reaction proceeding
in the opposite direction, the rate of NADH oxidation is indi-
cated by the rate of decrease in absorbance at 340 nm. Many
dehydrogenases can be directly assayed by this procedure. In
addition, the concentrations of a product formed in a nonox-
idative reaction can often be determined by oxidizing the
product in a dehydrogenase- NAD® system. Such a measure-
ment of concentrations of NAD® or NADH by their absorption
of ultraviolet light is used not only in the research laboratory
but also in many clinical analyses.

Wavelength (nm)

▲ Ultraviolet absorption spectra of NAD© and NADH.

10.10 Experimental Methods for Studying
Metabolism

The complexity of many metabolic pathways makes them difficult to study. Reaction
conditions used with isolated reactants in the test tube (in vitro ) are often very different
from the reaction conditions in the intact cell (in vivo). The study of the chemical events
of metabolism is one of the oldest branches of biochemistry, and many approaches have
been developed to characterize the enzymes, intermediates, flux, and regulation of
metabolic pathways.

A classical approach to unraveling metabolic pathways is to add a substrate to prepa-
rations of tissues, cells, or subcellular fractions and then follow the emergence of inter-
mediates and end products. The fate of a substrate is easier to trace when the substrate
has been specifically labeled. Since the advent of nuclear chemistry, isotopic tracers have
been used to map the transformations of metabolites. For example, compounds contain-
ing atoms of radioactive isotopes such as 3 H or 14 C can be added to cells or other prepa-
rations, and the radioactive compounds produced by anabolic or catabolic reactions can
be purified and identified. Nuclear magnetic resonance (NMR) spectroscopy can trace
the reactions of certain isotopes. It can also be employed to study the metabolism of
whole animals (including humans) and is being used for clinical analysis.

Verification of the steps of a particular pathway can be accomplished by reproduc-
ing the separate reactions in vitro using isolated substrates and enzymes. Individual
enzymes have been isolated for almost all known metabolic steps. By determining the
substrate specificity and kinetic properties of a purified enzyme, it is possible to draw
some conclusions regarding the regulatory role of that enzyme. This reductionist ap-
proach has led to many of the key concepts in this book. It’s the approach that allows us
to understand the relationship between structure and function. However, a complete as-
sessment of the regulation of a pathway requires analysis of metabolite concentrations
in the intact cell or organism under various conditions.

322

CHAPTER 10 Introduction to Metabolism

Valuable information can be acquired by studying mutations in single genes associ-
ated with the production of inactive or defective individual enzyme forms. Whereas
some mutations are lethal and not transmitted to subsequent generations, others can be
tolerated by the descendants. The investigation of mutant organisms has helped identify
enzymes and intermediates of numerous metabolic pathways. Typically, a defective en-
zyme results in a deficiency of its product and the accumulation of its substrate or a
product derived from the substrate by a branch pathway. This approach has been ex-
tremely successful in identifying metabolic pathways in simple organisms such as bacte-
ria, yeast, and Neurospora (Box 7.4). In humans, enzyme defects are manifested in meta-
bolic diseases. Hundreds of single-gene diseases are known. Some are extremely rare,
and others are fairly common; some are tragically severe. In cases where a metabolic
disorder produces only mild symptoms, it appears that the network of metabolic reac-
tions contains enough overlap and redundancy to allow near-normal development of
the organism.

In instances where natural mutations are not available, mutant organisms can be
generated by treatment with radiation or chemical mutagens (agents that cause muta-
tion). Biochemists have characterized entire pathways by producing a series of mutants,
isolating them, and examining their nutritional requirements and accumulated
metabolites. More recently, site-directed mutagenesis (Box 6.1) has proved valuable in
defining the roles of enzymes. Bacterial and yeast systems have been the most widely
used for introducing mutations because large numbers of these organisms can be
grown in a short period of time. It is possible to produce animal models — particularly
insects and nematodes — in which certain genes are not expressed. It is also possible to
delete certain genes in vertebrates. “Gene knockout” mice, for instance, provide an ex-
perimental system for investigating the complexities of mammalian metabolism.

In a similar fashion, investigating the actions of metabolic inhibitors has helped
identify individual steps in metabolic pathways. The inhibition of one step of a pathway
affects the entire pathway. Because the substrate of the inhibited enzyme accumulates, it
can be isolated and characterized more easily. Intermediates formed in steps preceding
the site of inhibition also accumulate. The use of inhibitory drugs not only helps in the
study of metabolism but also determines the mechanism of action of the drug, often
leading to improved drug variations.

Summary

1. The chemical reactions carried out by living cells are collectively
called metabolism. Sequences of reactions are called pathways.
Degradative (catabolic) and synthetic (anabolic) pathways pro-
ceed in discrete steps.

2. Metabolic pathways are regulated to allow an organism to re-
spond to changing demands. Individual enzymes are commonly
regulated by allosteric modulation or reversible covalent modifi-
cation.

3. The major catabolic pathways convert macromolecules to smaller,
energy-yielding metabolites. The energy released in catabolic reac-
tions is conserved in the form of ATP, GTP, and reduced coenzymes.

4. Within a cell or within a multicellular organism, metabolic processes
are sequestered.

5. Metabolic reactions are in a steady state. If the steady state con-
centration of reactants and products is close to the equilibrium
ratio the reaction is said to be a near-equilibrium reaction. If the
steady state concentrations are far from equilibrium the reaction
is said to be metabolically irreversible.

6. The actual free energy change (AG) of a reaction inside a cell
differs from the standard free energy change (A G°').

7. Hydrolytic cleavage of the phosphoanhydride groups of ATP re-
leases large amounts of free energy.

8. The energy of ATP is made available when a terminal phosphoryl
group or a nucleotidyl group is transferred. Some metabolites
with high phosphoryl group transfer potentials can transfer their
phosphoryl groups to ADP to produce ATP. Such metabolites are
called energy-rich compounds.

9. Thioesters, such as acyl coenzyme A, can donate acyl groups and
can sometimes also generate ATP equivalents.

10. The free energy of biological oxidation reactions can be captured
in the form of reduced coenzymes. This form of energy is meas-
ured as the difference in reduction potentials.

11. Metabolic pathways are studied by characterizing their enzymes,
intermediates, flux, and regulation.

Problems 323

Problems

1. A biosynthetic pathway proceeds from compound A to com-
pound E in four steps and then branches. One branch is a two-step
pathway to G, and the other is a three-step pathway to J. Substrate
A is a feed-forward activator of the enzyme that catalyzes the syn-
thesis of E. Products G and J are feedback inhibitors of the initial
enzyme in the common pathway, and they also inhibit the first
enzyme after the branch point in their own pathways.

(a) Draw a diagram showing the regulation of this metabolic
pathway.

(b) Why is it advantageous for each of the two products to in-
hibit two enzymes in the pathway?

2. Glucose degradation can be accomplished by a combination of
the glycolytic and citric acid pathways. The enzymes for glycolysis
are located in the cytosol, while the enzymes for the citric acid
cycle are located in the mitochondria. What are two advantages in
separating the enzymes for these major carbohydrate degradation
pathways in different cellular compartments?

3. In bacteria, the glycolytic and citric acid cycle pathways are both
cytosolic. Why don’t the “advantages” in Question 2 apply to
bacteria?

4. In multistep metabolic pathways, enzymes for successive steps
may be associated with each other in multienzyme complexes
or be bound in close proximity on membranes. Explain the
major advantage of having enzymes organized in either of these
associations.

5. (a) Calculate the K eq at 25°C and pH 7.0 for the following reaction

using the data in Table 10.4.

Glycerol 3-phosphate + H 2 0 — > glycerol + Pj.

(b) The final step in the pathway for the synthesis of glucose
from lactate (gluconeogenesis) is:

Glucose 6-P + H 2 0 — » glucose + Pj.

When glucose 6-P is incubated with the proper enzyme and
the reaction runs until equilibrium has been reached, the
final concentrations are found to be: glucose 6-P (0.035 mM),
glucose (100 mM), and Pi (100 mM). Calculate AG 0 ' at 25°C and
pH 7.0.

6. AG° for the hydrolysis of phosphoarginine is —32 kj mol -1 .

(a) What is the actual free energy change for the reaction at 25°C
and pH 7.0 in resting lobster muscle, where the concentra-
tions of phosphoarginine, arginine, and Pi are 6.8 mM,
2.6 mM, and 5 mM, respectively?

(b) Why does this value differ from AG 0 '?

(c) High-energy compounds have large negative free energies of
hydrolysis, indicating that their reactions with water proceed
almost to completion. How can millimolar concentrations of
acetyl Co A, whose AG 0 'hydrolysis is — 32 kj mol -1 , exist in cells?

7. Glycogen is synthesized from glucose- 1 -phosphate. Glucose- 1-
phosphate is activated by a reaction with UTP, forming UDP-glu-
cose and pyrophosphate (PPi).

Glucose-1 -phosphate + UTP — » UDP-glucose + PPj

UDP- glucose is the substrate for the enzyme glycogen synthase
which adds glucose molecules to the growing carbohydrate chain.
The AG 0 ' value for the condensation of UTP with glucose- 1-
phosphate to form UDP-glucose is approximately 0 kj mol -1 .

The PPi that is released is rapidly hydrolyzed by inorganic py-
rophosphatase. Determine the overall AG 0 ' value if the forma-
tion of UDP-glucose is coupled to the hydrolysis of PPi.

8. (a) A molecule of ATP is usually consumed within a minute after

synthesis, and the average human adult requires about 65 kg
of ATP per day. Since the human body contains only about
50 grams of ATP and ADP combined, how it is possible that
so much ATP can be utilized?

(b) Does ATP have a role in energy storage?

9. Phosphocreatine is produced from ATP and creatine in mam-
malian muscle cells at rest. What ATP/ADP ratio is necessary to
maintain a phosphocreatine/creatine ratio of 20:1? (To maintain
the coupled reaction at equilibrium, the actual free energy change
must be zero.)

10. Amino acids must be covalently attached to a ribose hydroxyl
group on the correct tRNA (transfer RNA) prior to recognition
and insertion into a growing polypeptide chain. The overall reac-
tion carried out by the amino acyl tRNA synthetase enzymes is:

Amino acid + HO-tRNA + ATP >

amino acyl-O-tRNA + AMP + 2Pj

Assuming this reaction proceeds through an acyl adenylate inter-
mediate, write all the steps involved in this enzyme- catalyzed
reaction.

11. When a mixture of glucose 6-phosphate and fructose 6-phosphate
is incubated with the enzyme phosphohexose isomerase, the final
mixture contains twice as much glucose 6-phosphate as fructose
6-phosphate. Calculate the value of AG 0 '.

Glucose 6-phosphate < — » fructose 6-phosphate

12. Coupling ATP hydrolysis to a thermodynamically unfavorable reac-
tion can markedly shift the equilibrium of the reaction.

(a) Calculate K eq for the energetically unfavorable biosynthetic
reaction A — » B when A G° = + 25 kj mol -1 at 25°C.

(b) Calculate K eq for the reaction A — » B when it is coupled to
the hydrolysis of ATP. Compare this value to the value in Part (a).

(c) Many cells maintain [ATP]/ [ADP] ratios of 400 or more. Cal-
culate the ratio of [B] to [A] when [ATP]: [ADP] is 400:1 and
[Pi] is constant at standard conditions. How does this ratio
compare to the ratio of [B] to [A] in the uncoupled reaction?

13. Using data from Table 10.5, write the coupled reaction that would
occur spontaneously for the following pairs of molecules under
standard conditions:

(a) Cytochrome /and cytochrome b 5

(b) Fumarate/succinate and ubiquinone/ubiquinol (Q/QH 2 )

14. Using data from Table 10.5, calculate the standard reduction po-
tential and the standard free energy change for each of the follow-
ing oxidation-reduction reactions:

(a) Ubiquinol (QH 2 ) + 2 cytochrome c (Fe^)

ubiquinone (Q) + 2 cytochrome c (Fe®) + 2 H©

(b) Succinate + y 2 0 2 fumarate + H 2 0

324 CHAPTER 10 Introduction to Metabolism

15 . Lactate dehydrogenase is an NAD-dependent enzyme that cat-
alyzes the reversible oxidation of lactate.

coo e

HO— C — H

ch 3

NAD© NADH, H©

COO©

c = o
I

ch 3

16 . Using the standard reduction potentials for Q and FAD in Table 10.5,
show that the oxidation of FADH 2 by Q liberates enough energy to
drive the synthesis of ATP from ADP and Pj under cellular conditions
where [FADH 2 ] = 5 mM, [FAD] = 0.2 mM, [Q] = 0.1 mM,
and [QH 2 ] = 0.05 mM. Assume that AG for ATP synthesis from
ADP and Pj is +30 kj mol -1 .

Initial reaction rates are followed spectrophotometrically at 340 nm
after addition of lactate, NAD©, lactate dehydrogenase, and
buffer to the reaction vessel. When the change in absorbance at
340 nm is monitored over time, which graph is representative of
the expected results? Explain.

Time

■xt

Time

Selected Readings

Alberty, R. A. (1996). Recommendations
for nomenclature and tables in biochemical
thermodynamics. Eur. J. Biochem.

240:1-14.

Alberty, R. A. (2000). Calculating apparent equi-
librium constants of enzyme -catalyzed reactions
at pH 7. Biochem. Educ. 28:12-17.

Burbaum, J. J., Raines, R. T., Albery, W. J., and
Knowles, J. R. (1989). Evolutionary optimization of
the catalytic effectiveness of an enzyme. Biochem.
28:9293-9305.

Edwards, R. A. (2001). The free energies of meta-
bolic reactions (AG) are not positive. Biochem.
Mol. Bio. Educ. 29:101-103.

Hayes, D. M., Kenyon, G. L., and Kollman, P. A.
(1978). Theoretical calculations of the hydrolysis
energies of some “high-energy” molecules. 2. A
survey of some biologically important hydrolytic
reactions. /. Am. Chem. Soc. 100:4331-4340.

Schmidt. S., Sunyaev, S., Bork. P., and Dandekar, T.
(2003). Metabolites: a helping hand for pathway
evolution? Trends Biochem. Sci. 28:336-341.

Silverstein, T. (2005). Redox redox: a response to
Feinman’s “Oxidation-reduction calculations in
the biochemistry course.” Biochem. Mol. Bio. Educ.
33:252-253.

Tohge, T., Nunes-Nesi, A., and Fernie, A. R. (2009).
Finding the paths: metabolomics and approaches
to metabolic flux analysis. The Biochem. Soc. (June
2009):8-12.

Yus, E., et al. (2009). Impact of genome reduction
on bacterial metabolism and its regulation. Science
326:1263-1272.

o c

_ o

° o o o

° o

o o

° c

o o

T he first three metabolic pathways we examine are central to both carbohydrate
metabolism and energy generation. Gluconeogenesis is the main pathway for
synthesis of hexoses from three carbon precursors. As the name of the pathway
indicates, glucose is the primary end product of gluconeogenesis. This biosynthetic
pathway will be described in the next chapter. Glucose, and other hexoses, can be the
precursors for synthesis of many complex carbohydrates. Glucose can also be degraded
in a catabolic glycolytic pathway with recovery of the energy used in its synthesis. In
glycolysis, the subject of this chapter, glucose is converted to the three-carbon acid
pyruvate. Pyruvate has several possible fates, one of which is oxidative decarboxylation
to form acetyl CoA. The third pathway is the citric acid cycle, described in Chapter 13.
This is the route by which the acetyl group of acetyl CoA is oxidized to carbon dioxide
and water. One of the important intermediates in the citric acid cycle, oxaloacetate, is
also an intermediate in the synthesis of glucose from pyruvate. Figure 11.1 shows the re-
lationship among the three pathways. All three pathways play a role in the formation
and degradation of noncarbohydrate molecules such as amino acids and lipids.

We present the reactions of glycolysis, gluconeogenesis, and the citric acid cycle in
more detail than those of other metabolic pathways in this book but the same principles
apply to all pathways. We introduce many biomolecules and enzymes, some of which
appear in more than one pathway. Keep in mind that the chemical structures of the
metabolites prompt the enzyme names and that the names of the enzymes reflect the
substrate specificity and the type of reaction catalyzed. A confident grasp of terminol-
ogy will prepare you to enjoy the chemical elegance of metabolism. However, do not
lose sight of the major concepts and general strategies of metabolism while memorizing
the details. The names of particular enzymes might fade over time but we hope you will
retain an understanding of the patterns and purposes behind the interconversion of
metabolites in cells.

In this book we follow the tradition of presenting glycolysis as our first metabolic
pathway. The catabolism of glucose is a major source of energy in animals. The details
of the various reactions, and their regulation, are well known.

The glycolytic sequence of reactions
is perhaps the best understood and
most studied multi-enzyme system
of the cell. The pattern of interplay
between enzymes and substrates in
this relatively simple multi-enzyme
system applies to all the multi-
enzyme systems of the cell, espe-
cially the very complex systems
involved in respiration and
photosynthesis.

— Albert Lehninger (1965),
Bioenergetics , p. 75

Top: Wine, beer and bread. For centuries, wineries, breweries, and bakeries have exploited the basic biochemical pathway
of glycolysis where glucose is converted to ethanol and C0 2 .

325

326

CHAPTER 11 Glycolysis

Gluconeogenesis

Glycolysis

Phosphoenolpyruvate

▲ Figure 11.1

Gluconeogenesis, glycolysis, and the citric
acid cycle. Glucose is synthesized from
pyruvate via oxaloacetate and phospho-
enolpyruvate. In glycolysis, glucose is de-
graded to pyruvate. Many (but not all) of the
steps in glycolysis are the reverse of the glu-
coneogenesis reactions. The acetyl group of
pyruvate is transferred to coenzyme A (CoA)
and oxidized to carbon dioxide by the citric
acid cycle. Energy in the form of ATP equiv-
alents is required for the synthesis of glu-
cose. Some of this energy is recovered in
glycolysis but much more is recovered as a
result of the citric acid cycle.

KEY CONCEPT

The main energy gain in glycolysis is due
to production of NADH molecules.

11.1 The Enzymatic Reactions of Glycolysis

Glycolysis is a sequence of ten enzyme -catalyzed reactions by which glucose is con-
verted to pyruvate (Figure 11.2 on page 328). The conversion of one molecule of glu-
cose to two molecules of pyruvate is accompanied by the net conversion of two mole-
cules of ADP to two molecules of ATP and the reduction of two molecules of NADH®
to two molecules NADH. The enzymes of this pathway are found in most living species
and are located in the cytosol. The glycolytic pathway is active in all differentiated cell
types in multicellular organisms. In some mammalian cells (such as those in the retina
and some brain cells), it is the only ATP-producing pathway.

The net reaction of glycolysis is shown in Reaction 11.1.

Glucose + 2 ADP + 2 NAD® + 2 P; ->

2 Pyruvate + 2 ATP + 2 NADH + 2 H® + 2 H 2 Q (11.1)

The ten reactions of glycolysis are listed in Table 11.1. They can be divided into two
stages: the hexose stage and the triose stage. The left page of Figure 1 1.2 shows the hex-
ose stage. At Step 4, the C-3 — C-4 bond of the hexose is cleaved to produce two trioses.
From that point on the intermediates of the pathway are triose phosphates. Two triose
phosphates are formed from fructose 1,6-frisphosphate. Dihydroxyacetone phosphate is
converted to glyceraldehyde 3 -phosphate in Step 5 and glyceraldehyde 3 -phosphate
continues through the pathway. All subsequent steps of the triose stage of glycolysis
(right page of Figure 11.2) are traversed by two molecules for each molecule of glucose
metabolized.

Two molecules of ATP are converted to ADP in the hexose stage of glycolysis. In the
triose stage, four molecules of ATP are formed from ADP for each molecule of glucose me-
tabolized. Thus, glycolysis has a net yield of two molecules of ATP per molecule of glucose.

ATP consumed per glucose: 2 (hexose stage)

ATP produced per glucose: 4 (triose stage) (11.2)

Net ATP production per glucose: 2

The first and third reactions of glycolysis are coupled to the utilization of ATP.
These priming reactions help drive the pathway in the direction of glycolysis since the
reverse reactions are thermodynamically favored in the absence of ATP. Two later inter-
mediates of glycolysis have sufficient group transfer potentials to allow the transfer of a
phosphoryl group to ADP producing ATP (Steps 7 and 10). Step 6 is coupled to the syn-
thesis of reducing equivalents in the form of NADH. Each molecule of NADH is equiv-
alent to several molecules of ATP (Section 10.9) so the net energy gain in glycolysis is
mostly due to production of NADH.

11.2 The Ten Steps of Glycolysis

Now we examine the chemistry and enzymes of each glycolytic reaction. As you read, pay
attention to the chemical logic and economy of the pathway. Consider how each chemical
reaction prepares a substrate for the next step in the process. Note, for example, that a
cleavage reaction converts a hexose to two trioses, not to a two -carbon compound and a
tetrose. The two trioses rapidly interconvert allowing both products of the cleavage reac-
tion to be further metabolized by the action of one set of enzymes, not two. Finally, be
aware of how ATP is both consumed and produced in glycolysis. We have already seen a
number of examples of the transfer of the chemical potential energy of ATP (e.g., in Sec-
tion 10.7) but the reactions in this chapter are our first detailed examples of how the en-
ergy released by oxidation reactions is captured for use in other biochemical pathways.

1. Hexokinase

In the first reaction of glycolysis, the y -phosphoryl group of ATP is transferred to the
oxygen atom at C-6 of glucose producing glucose 6-phosphate and ADP (Figure 1 1.3 on

1 1 .2 The Ten Steps of Glycolysis 327

Table 11.1 The reactions and enzymes of glycolysis

1. Glucose + ATP > Glucose 6-phosphate + ADP + H©

Hexokinase, glucokinase

2. Glucose 6-phosphate Fructose 6-phosphate

Glucose-6-phosphate isomerase

3. Fructose 6-phosphate + ATP > Fructose 1,6-b/sphosphate + ADP + H©

Phosphofructokinase-1

4. Fructose 1,6-b/sphosphate Dihydroxyacetone phosphate + Glyceraldehyde 3-phosphate

Aldolase

5. Dihydroxyacetone phosphate Glyceraldehyde 3-phosphate

6. Glyceraldehyde 3-phosphate + NAD© + Pj 1,3-8/sphosphoglycerate + NADH + H©

7. 1,3-£/sphosphoglycerate + ADP 3-Phosphoglycerate + ATP

Triose phosphate isomerase
Glyceraldehyde 3-phosphate dehydrogenase
Phosphoglycerate kinase

8. 3-Phosphoglycerate 2-Phosphoglycerate

Phosphoglycerate mutase

9. 2-Phosphoglycerate Phosphoenolpyruvate + H 2 0

Enolase

10. Phosphoenolpyruvate + ADP + H© > Pyruvate + ATP

Pyruvate kinase

page 330). This phosphoryl group transfer reaction is catalyzed by hexokinase. Kinases
catalyze four reactions in the glycolytic pathway — Steps 1, 3, 7, and 10.

The hexokinase reaction is regulated making it a metabolically irreversible reac-
tion. Cells need to maintain a relatively high concentration of glucose 6-phosphate and
a low internal concentration of glucose. As we’ll see in Section 1 1.5B, the reverse reaction
is inhibited by glucose 6-phosphate. Hexokinases from yeast and mammalian tissues
have been thoroughly studied. These enzymes have a broad substrate specificity; they
catalyze the phosphorylation of glucose and mannose, and of fructose when it is present
at high concentrations.

Multiple forms, or isozymes, of hexokinase occur in many eukaryotic cells.
(Isozymes are different proteins from one species that catalyze the same chemical reac-
tion.) Four hexokinase isozymes have been isolated from mammalian liver. All four are
found in varying proportions in other mammalian tissues. These isozymes catalyze the
same reaction but have different K m values for glucose. Hexokinases I, II, and III have
K m values of about 10 -6 to 10 -4 M, whereas hexokinase IV, also called glucokinase, has a
much higher K m value for glucose (about 10 -2 M). In eukaryotes, glucose is taken up
and secreted by passive transport using various glucose transporters (GLUT). The con-
centration of glucose in the blood and the cell cytoplasm is usually below the K m of glu-
cokinase for glucose. At these low concentrations the other hexokinase isozymes catalyze
the phosphorylation of glucose. With high glucose levels, glucokinase is active. Because
glucokinase is never saturated with glucose, the liver can respond to large increases in
blood glucose by phosphorylating it for entry into glycolysis or the glycogen synthesis
pathway.

In most bacteria, the uptake of glucose is coupled to the phosphorylation of glu-
cose to glucose 6-phosphate via the phosphoenolpyruvate sugar transport system (Sec-
tion 21.7B). The phosphoryl group is donated by phosphoenolpyruvate. Hexokinases
and glucokinases can be found in bacteria but they play a minor role in glycolysis be-
cause, unlike the situation in eukaryotic cells, the bacterial enzymes rarely encounter
free glucose in their cytoplasm.

2. Glucose 6-Phosphate Isomerase

In the second step of glycolysis, glucose 6-phosphate isomerase catalyzes the conversion of
glucose 6-phosphate (an aldose) to fructose 6-phosphate (a ketose), as shown in Figure
1 1.4. The enzyme is also known as phosphoglucose isomerase (PGI). Isomerases intercon-
vert aldoses and ketoses that have identical configurations at all other chiral atoms.

The a anomer of glucose 6-phosphate (a-D-glucopyranose 6-phosphate) preferen-
tially binds to glucose 6-phosphate isomerase. The open-chain form of glucose 6-phos-
phate is then generated within the active site of the enzyme, and an aldose-to-ketose con-
version occurs. The open-chain form of fructose 6-phosphate cyclizes to form
a-D-fructofuranose 6-phosphate. The mechanism of glucose 6-phosphate isomerase is
similar to the mechanism of triose phosphate isomerase (Section 6.4A).

Glucose 6 -phosphate isomerase is highly stereospecific. For example, in the reverse
reaction catalyzed by this enzyme fructose 6-phosphate (in which C-2 is not chiral) is

328 CHAPTER 11 Glycolysis

Figure 1 1.2 ►

Conversion of glucose to pyruvate by glycolysis. At Step 4, the
hexose molecule is split in two, and the remaining reactions
of glycolysis are traversed by two triose molecules. ATP is
consumed in the hexose stage and generated in the triose
stage.

Transfer of a phosphoryl Hexokinase, glucokinase

group from ATP to glucose

ATP

ADP + H®

Isomerization

(5) Glucose 6-phosphate isomerase

Fructose 6-phosphate

Transfer of a second

phosphoryl group from ATP (3) Phosphofructokinase-1
to fructose 6-phosphate

ATP

ADP + H®

».©

Fructose 1,6-b/sphosphate

C-3 — C-4 bond
cleavage , yielding
two triose phosphates

(4) Aldolase

C = 0 H — C — OH

1 1 ©

CH 2 OH CH 2 0P0 3 (i)

Dihydroxyacetone phosphate Glyceraldehyde 3-phosphate

1 1 .2 The Ten Steps of Glycolysis 329

O H

% /

CH 2 OH C

c=0 > H — c — OH Those phosphate ^

isomerase

ch 2 opo 3 ® ch 2 opo 3 ©

Dihydroxyacetone phosphate Glyceraldehyde 3-phosphate

NAD®+P|
NADH + H©

Glyceraldehyde
3-phosphate @
dehydrogenase

O x OP0 3 ©

H — C — OH

ch 2 opo 3 ©

1 ,3-£/sphosphoglycerate

ADP

ATP

Phosphoglycerate kinase ( 7 )

coo©

H — C — OH

CH 2 OPO^

3-Phosphoglycerate

Phosphoglycerate mutase (§)

COO 0

H — C — 0P0 3 ®
CH 2 OH

2-Phosphoglycerate

H 2 0

COO©

C— OPO 3 ©

ch 2

Phosphoenolpyruvate

^ ADP + H®

v » ATP

Enolase

Pyruvate kinase

coo©

c =0

ch 3

Pyruvate

Rapid interconversion of
triose phosphates

Oxidation and phosphorylation ,
yielding a high-energy
mixed -acid anhydride

Transfer of a high-energy
phosphoryl group to ADP ’
yielding ATP

Intramolecular

phosphoryl-group

transfer

Dehydration to
an energy-rich enol ester

Transfer of a high-energy
phosphoryl group to ADP,
yielding ATP

330 CHAPTER 11 Glycolysis

0 CT

0 O — P — O — P — O — P — O — Adenosine

II \J II II

OH O O O

MgATP©

Hexokinase

©o" M9 '"o©

^ I I

0 O— P = 0 0 O— P — O— P — O — Adenosine

I II II

0 0 0

MgADP 0

+ H

Glucose 6-phosphate

▲ Figure 1 1.3

Phosphoryl group transfer reaction catalyzed
by hexokinase. This reaction occurs by at-
tack of the C-6 hydroxyl oxygen of glucose
on the y-phosphorus of MgATP 0 . MgADP®
is displaced, and glucose 6-phosphate is
generated. All four kinases in glycolysis cat-
alyze direct nucleophilic attack of a hydroxyl
group on the terminal phosphoryl group of
ATP (and/or its reverse under cellular condi-
tions). (Mg©, shown explicitly here, is also
required in the other kinase reactions in this
chapter, although it is not shown for those
reactions.)

The hexokinase mechanism is a classic
example of induced fit (Section 6.5C).

We discuss the regulation of glycolysis
in detail in Section 1 1.5.

We explore glycogen synthesis in
Section 12.5.

converted almost exclusively to glucose 6-phosphate. Only traces of mannose 6-phos-
phate, the C-2 epimer of glucose 6-phosphate, are formed.

The glucose 6-phosphate isomerase reaction is a near-equilibrium reaction. The re-
verse reaction is part of the pathway for the biosynthesis of glucose.

3. Phosphofructokinase-1

Phosphofructokinase-l (PFK-1) catalyzes the transfer of a phosphoryl group from
ATP to the C-l hydroxyl group of fructose 6-phosphate producing fructose
1,6-frisphosphate. The “bis” in frzsphosphate indicates that the two phosphoryl groups
are attached to different carbon atoms (cf. diphosphate).

Fructose 6-phosphate Fructose 1,6-b/sphosphate

(11.3)

Note that the reaction catalyzed by glucose 6-phosphate isomerase produces a-D-fruc-
tose 6-phosphate. However, it is the /3-d anomer that is the substrate for the next step in
glycolysis — the one catalyzed by phosphofructokinase-1. The a and /3 anomers of fruc-
tose 6-phosphate equilibrate spontaneously (Section 8.2). This interconversion is
extremely rapid in aqueous solution and has no effect on the overall rate of glycolysis.

The reaction catalyzed by PFK- 1 is metabolically irreversible indicating that the
activity of the enzyme is regulated. In fact, this step is a critical control point for the reg-
ulation of glycolysis in most species. The PFK-1 catalyzed reaction is the first committed
step of glycolysis because some substrates other than glucose can enter the glycolytic
pathway by direct conversion to fructose 6-phosphate, thus bypassing the steps cat-
alyzed by hexokinase and glucose 6-phosphate isomerase (Section 11. 6C). (The meta-
bolically irreversible reaction catalyzed by hexokinase is not the first committed step.)
Another reason for regulating PFK- 1 activity has to do with the competing glycolysis
and gluconeogenesis pathways (Figure 11.1). PFK-1 activity must be inhibited when
glucose is being synthesized.

PFK-1 is one of the classic allosteric enzymes. Recall that the bacterial enzyme is ac-
tivated by ADP and allosterically inhibited by phosphoenolpyruvate (Section 5.10A).
The activity of the mammalian enzyme is regulated by AMP and citrate (Section 1 1.6C).

PFK-1 has the suffix “1” because there is a second phosphofructokinase that cat-
alyzes the synthesis of fructose 2,6-Hsphosphate instead of fructose l,6-Z?isphosphate.
This second enzyme, which we will encounter later in this chapter, is known as PFK- 2.

4. Aldolase

The first three steps of glycolysis prepare the hexose for cleavage into two triose phos-
phates, glyceraldehyde 3 -phosphate and dihydroxyacetone phosphate.

1 1 .2 The Ten Steps of Glycolysis 331

Glucose 6-phosphate
(u-D-glucopyranose form)

H /°

1 ch 2 oh

— C — OH

-u

2 1

Glucose

2 I

6-phosphate
isomerase
>

HO — C — H

3 |

-C — OH

H — C— OH

4 1

-u-

H — C— OH

b |

6 ch 2 opo 3 ©

6 ch 2 opo :

Glucose 6-phosphate
(open-chain form)

Fructose 6-phosphate
(open-chain form)

Fructose 6-phosphate
(a-D-fructofuranose form)

▲ Figure 1 1.4

Conversion of glucose 6-phosphate to fructose 6-phosphate. This aldose-ketose isomerization is
catalyzed by glucose 6-phosphate isomerase.

Dihydroxyacetone phosphate (DHAP) is derived from C-l to C-3 of fructose
1,6-fcphosphate, and glyceraldehyde 3-phosphate (GAP) is derived from C-4 to C-6. The
enzyme that catalyzes the cleavage reaction is fructose 1,6-frisphosphate aldolase, com-
monly shortened to aldolase. Aldol cleavage is a common mechanism for cleaving C — C
bonds in biological systems and for C — C bond formation in the reverse direction.

BOX 1 1 .1 A BRIEF HISTORY OF THE GLYCOLYTIC PATHWAY

Glycolysis was one of the first metabolic pathways to be eluci-
dated. It played an important role in the development of bio-
chemistry. In 1897, Eduard Buchner (Section 1.1) discovered
that bubbles of carbon dioxide were released from a mixture
of sucrose and a cell- free yeast extract. He concluded that fer-
mentation was occurring in his cell- free extract. More than 20
years earlier, Louis Pasteur had shown that yeast cells ferment
sugar to alcohol (i.e., produce ethanol and C0 2 ) but Buchner
showed that intact cells were not required. Buchner named
the fermenting activity zymase. Today, we recognize that the
zymase of yeast extracts is not a single enzyme but a mixture
of enzymes that together catalyze the reactions of glycolysis.

The steps of the glycolytic pathway were gradually dis-
covered by analyzing the reactions catalyzed by extracts of

yeast or muscle. In 1905, Arthur Harden and William John
Young found that when the rate of glucose fermentation by
yeast extract decreased it could be restored by adding inor-
ganic phosphate. Harden and Young assumed that phosphate
derivatives of glucose were being formed. They succeeded in
isolating fructose 1,6-frzsphosphate and showed that it is an
intermediate in the fermentation of glucose because it too is
fermented by cell-free yeast extracts. Harden was awarded the
Nobel Prize in Chemistry in 1929 for his work on glycolysis.

By the 1940s, the complete glycolytic pathway in eukary-
otes — including its enzymes, intermediates, and coenzymes —
was known. The further characterization of individual enzymes
and studies of the regulation of glycolysis and its integration
with other pathways have taken many more years. In bacteria,
the classic glycolytic pathway is called the Embden-Meyerhof-
Parnas pathway after Gustav Embden (1874-1933), Otto
Meyerhof (1884-1951), and Jacob Parnas (1884-1949). The
bacterial pathway differs in some minor ways from the
eukaryotic pathway. In 1922 Meyerhof was awarded the Nobel
Prize in Physiology or Medicine for his work on the production
of lactic acid in muscle cells.

▲ Louis Pasteur (1822-1895).

a Arthur Harden (1865-1940).

332

CHAPTER 11 Glycolysis

2 I = °

H y O

HO— 3 C — H

(1) ch 2 opo 3 ©

(4) ,

Aldolase

H— 4 C— OH < >

4 |

H^C-OH

H — C — OH

(3)CH 2 OH

(6) ch 2 opo 3 <

6 ch 2 opo 3 ©

Fructose 1,6-b/sphosphate

Dihydroxyacetone

Glyceraldehyde

phosphate

3-phosphate

(11>

T Figure 1 1.5

Mechanism of aldol cleavage catalyzed by
aldolases. Fructose 1,6-b/sphosphate is the
aldol substrate. Aldolases have an electron-
withdrawing group ( — X) that polarizes the
C-2 carbonyl group of the substrate. Class I
aldolases use the amino group of a lysine
residue at the active site, and the other
class II aldolases use Zn© for this purpose.
A basic residue (designated — B:) removes a
proton from the C-4 hydroxyl group of the
substrate.

There are two distinct classes of aldolases, class I enzymes are found in plants and
animals; class II enzymes are more common in bacteria, fungi, and protists. Many
species have both types of enzyme. Class I and class II aldolases are unrelated. The en-
zymes have very different structures and sequences in spite of the fact that they catalyze
the same reaction. This is an example of convergent evolution.

The two classes of aldolase have slightly different mechanisms. Class I aldolases in-
volve formation of a covalent Schiff base between lysine and pyruvate derivatives (Sec-
tion 6.3) and class II aldolases use a metal ion cofactor (Figures 1 1.5 and 1 1.6).

The standard Gibbs free energy change for this reaction is strongly positive
(A G°' = +28 kj mol -1 ). Nevertheless, the aldolase reaction is a near-equilibrium reac-
tion (actual AG = 0) in cells where glycolysis is an important catabolic pathway. This
means that the concentration of fructose 1,6-Hsphosphate is very high relative to the
two trioses. (But see Problem 10).

The key to understanding the strategy of glycolysis lies in appreciating the signifi-
cance of the aldolase reaction. Its best to think of this as a near- equilibrium biosynthesis
reaction and not a degradation reaction. Aldolases evolved originally as enzymes that
could catalyze the synthesis of fructose 1,6-Hsphosphate. This reaction occurred at the
end of a biosynthesis pathway leading from pyruvate to glyceraldehyde 3 -phosphate
and dihydroxyacetone phosphate.

During glycolysis, flux in the triose stage is in the opposite direction — toward
pyruvate synthesis. The first steps of glycolysis — the hexose stage — are directed toward
formation of fructose 1,6-frisphosphate so that it can serve as substrate for the reversal
of the pathway leading to its synthesis. Keep in mind that the glucose biosynthesis path-
way (gluconeogenesis) evolved first. It was only after glucose became readily available
that pathways for its degradation evolved.

5. Triose Phosphate Isomerase

Of the two molecules produced by the splitting of fructose 1,6-Hsphosphate, only glyc-
eraldehyde 3 -phosphate is a substrate for the next reaction in the glycolytic pathway.

H ,0

(4) |

H “gjC OH

( 6 )

ch 2 opo

Glyceraldehyde

3-phosphate

ch 2 opo 3 ®

(2)

c=o

(3)

CH.OH

Dihydroxyacetone

phosphate

1 1 .2 The Ten Steps of Glycolysis 333

◄ Figure 11.6

Schiff base in the active site of aldolase. A

Schiff base forms between Lys-229 and di-
hydroxyacetone during the reaction catalyzed
by aldolase. Modified after St-Jean et al.
(2009). (Hydrogen atoms not shown.)

[PEB 3DF0]

The other product, dihydroxyacetone phosphate, is converted to glyceraldehyde 3 -phos-
phate in a near- equilibrium reaction catalyzed by triose phosphate isomerase.

CH 2 OH

c = o

ch 2 opo 3 ©

Triose

phosphate

isomerase

H /O

H — C— OH
CH 2 0P0 3 ®

Dihydroxyacetone

phosphate

Glyceraldehyde

3-phosphate

(11.5)

As glyceraldehyde 3 -phosphate is consumed in Step 6, its steady state concentration is
maintained by flux from dihydroxyacetone phosphate. In this way, two molecules of
glyceraldehyde 3 -phosphate are supplied to glycolysis for each molecule of fructose
1,6-frisphosphate split. Triose phosphate isomerase catalyzes a stereospecific reaction so
that only the D isomer of glyceraldehyde 3 -phosphate is formed.

Triose phosphate isomerase, like glucose 6-phosphate isomerase, catalyzes an
aldose-to-ketose conversion. The mechanism of the triose phosphate isomerase reaction
is described in Section 6.4A. The catalytic mechanisms of aldose-ketose isomerases
have been studied extensively, and the formation of an enzyme-bound enediolate inter-
mediate appears to be a common feature.

The fate of the individual carbon atoms of a molecule of glucose is shown in
Figure 11.7. This distribution has been confirmed by radioisotopic tracer studies in a
variety of organisms. Note that carbons 1, 2, and 3 of one molecule of glyceraldehyde
3 -phosphate are derived from carbons 4, 5, and 6 of glucose, whereas carbons 1,2, and 3
of the second molecule of glyceraldehyde 3-phosphate (converted from dihydroxyace-
tone phosphate) originate as carbons 3, 2, and 1 of glucose. When these molecules of
glyceraldehyde 3 -phosphate mix to form a single pool of metabolites, a carbon atom
from C-l of glucose can no longer be distinguished from a carbon atom from C-6 of
glucose.

The rate of the triose phosphate
isomerase reaction is close to the
theoretical limit for a diffusion
controlled reaction.

6. Glyceraldehyde 3-Phosphate Dehydrogenase

The recovery of energy from triose phosphates begins with the reaction catalyzed by
glyceraldehyde 3 -phosphate dehydrogenase. In this step, glyceraldehyde 3 -phosphate is
oxidized and phosphorylated to produce 1,3-Hsphosphoglycerate.

334

CHAPTER 11 Glycolysis

H — C— OH + NAD
CH 2 0P0 3 ©

+ Pi

Glyceraldehyde
3-phosphate
dehydrogenase
>

C> OP0 3 ©

H — C— OH + NADH + H®

I rr.

CH 2 0P0 3 © (11.6)

Glyceraldehyde 1,3-£/sphosphoglycerate

3-phosphate

This is an oxidation-reduction reaction; the oxidation of glyceraldehyde 3 -phosphate is
coupled to the reduction of NAD® to NADH. In some species the coenzyme is NADP® .

The oxidation of the aldehyde group of glyceraldehyde 3 -phosphate proceeds with a
large negative Gibbs standard free energy change, and some of this energy is conserved in
the acid-anhydride linkage of 1,3-fcphosphoglycerate. In the next step of glycolysis, the
C-l phosphoryl group of l,3-frzsphosphoglycerate is transferred to ADP to form ATP. The
remaining energy is conserved in the form of reducing equivalents (NADH). As we saw in
the previous chapter, each molecule of NADH is equivalent to several molecules of ATP.
Thus, this step of glycolysis is the main energy-producing step in the entire pathway.

The overall standard Gibbs free energy change (oxidation of the aldehyde and
reduction of NAD®) for this reaction is positive (AG o/ = +6.7 kj mol -1 ), which means
that the 1,3-Hsphosphate concentration should be much lower than that of glyceralde-
hyde 3 -phosphate at the near-equilibium conditions that exist inside the cell. However,
glyceraldehyde 3 -phosphate dehydrogenase associates with the next enzyme in the
pathway (phosphoglycerate kinase), to form a complex. The product of the first reaction,
1,3-frisphosphoglycerate, appears to be channeled directly into the active site of phospo-
glycerate kinase. In this way the two reactions are effectively linked to form a single reaction
and the effective concentration of 1,3-Hsphosphoglycerate is close to zero.

The NADH formed in the glyceraldehyde 3 -phosphate dehydrogenase reaction is
reoxidized, either by the membrane- associated electron transport chain (Chapter 14) or
in other reactions where NADH serves as a reducing agent, such as the reduction of
acetaldehyde to ethanol or of pyruvate to lactate (Section 1 1.3B). The concentration of
NAD® in most cells is low. Thus, it is essential to replenish it by reoxidizing NADH or
glycolysis will stop at this step. We will see in Section 1 1.3 that there are several different
ways of accomplishing this goal.

T Figure 1 1.7

Fate of carbon atoms from the hexose stage to
the triose stage of glycolysis. All numbers
refer to the carbon atoms in the original glu-
cose molecule.

H .0

1 ch 2 opo 3 ©

H — C — OH

2 1

-u-

r\j

HO— C — H

J | > _

HO — C — H

^ ^ J |

H-C-OH

H — C — OH

b |

H — C— OH

b |

6 ch 2 oh

6 ch 2 opo 3 ©

Glucose

Fructose

1,6-b/sphosphate

Aldolase/

XhUOH

* (2 fr O

Triose
phosphate
isomerase
>

(1) ch 2 opo 3©

Dihydroxyacetone

phosphate

H y O

( 3 ) ,

H^)C-

-OH

,CH,OPO

Glyceraldehyde

3-phosphate

H —rX. — OH

(6) ch 2 opo 3 ©

Glyceraldehyde

3-phosphate

1 1 .2 The Ten Steps of Glycolysis 335

7. Phosphoglycerate Kinase

Phosphoglycerate kinase catalyzes phosphoryl group transfer from the “high-energy”
mixed anhydride 1,3-Hsphosphoglycerate to ADP, generating ATP and 3 -phosphoglyc-
erate. The enzyme is called a kinase because of the reverse reaction in which 3 -phospho-
glycerate is phosphorylated.

C> OP0 3 ®

H — C— OH + ADP
CH 2 OP0 3 ®

1,3-B/sphosphoglycerate

(11.7)

COO

,©

Phosphoglycerate

kinase

H — C— OH + ATP

ch 2 opo 3 ©

3-Phosphoglycerate

Steps 6 and 7 together couple the oxidation of an aldehyde to a carboxylic acid with the
phosphorylation of ADP to ATP and the formation of a reducing equivalent.

Glyceraldehyde 3-phosphate + NAD® > 1,3-8/sphosphoglycerate + NADH + H®

1 ,3-8/sphosphoglycerate + ADP > 3-Phosphoglycerate + ATP

Glyceraldehyde 3-phosphate + NAD® + Pj + ADP > 3-Phosphoglycerate + NADH + H® + ATP

( 11 . 8 )

BOX 1 1.2 FORMATION OF 2,3-0/SPHOSPHOGLYCERATE IN RED BLOOD CELLS

An important function of glycolysis in red blood cells is the
production of 2,3-frzsphosphoglycerate, an allosteric in-
hibitor of the oxygenation of hemoglobin (Section 4.13C).
This metabolite is a reaction intermediate and cofactor in
Step 8 of glycolysis.

Erythrocytes contain Hsphosphoglycerate mutase. This
enzyme catalyzes the transfer of a phosphoryl group
from C-l to C-2 of 1,3 -^^phosphoglycerate, to form

2.3- Hsphosphoglycerate. As shown in the reaction scheme,

2.3- frisphosphoglycerate phosphatase catalyzes the hydrolysis
of excess 2,3BPG to 3 -phosphoglycerate, which can reenter
glycolysis and be converted to pyruvate.

The shunting of 1,3-frisphosphoglycerate through these
two enzymes bypasses phosphoglycerate kinase, which cat-
alyzes Step 7 of glycolysis, one of the two ATP- generating
steps. However, only a portion of the glycolytic flux in red
blood cells — about 20% — is diverted through the mutase
and phosphatase. Accumulation of free 2,3BPG (i.e., 2,3BPG
not bound to hemoglobin) inhibits fcphosphoglycerate mu-
tase. In exchange for diminished ATP generation, this bypass
provides a regulated supply of 2,3BPG, which is necessary for
the efficient release of 0 2 from oxyhemoglobin.

3-Phosphoglycerate

▲ Formation of 2,3-Z;/sphosphoglycerate (2,3BPG) in red blood cells.

336 CHAPTER 11 Glycolysis

BOX 1 1.3 ARSENATE POISONING

Arsenic, like phosphorus, is in Group V of the periodic table.
Arsenate (As0 4 ©) therefore, is an analog of inorganic phos-
phate. Arsenate competes with phosphate for its binding site
in glyceraldehyde 3 -phosphate dehydrogenase. Like phos-
phate, arsenate cleaves the energy-rich thioacyl-enzyme in-
termediate. However, arsenate produces an unstable analog
of 1,3-Hsphosphoglycerate, called l-arseno-3-phosphoglyc-
erate, which is rapidly hydrolyzed on contact with water. This
nonenzymatic hydrolysis produces 3-phosphoglycerate and
regenerates inorganic arsenate, which can again react with a
thioacyl-enzyme intermediate. In the presence of arsenate,
glycolysis can proceed from 3-phosphoglycerate, but the
ATP-producing reaction involving 1,3-frisphosphoglycerate is
bypassed. As a result, there is no net formation of ATP from
glycolysis. Arsenate is a poison because it can replace phos-
phate in many phosphoryl transfer reactions.

C> O— As — O e

f o 0

H — C— OH
CH 2 OP0 3 ©

1 -Arseno-3-phosphoglycerate

H 2 0 As0 4 ©

nonenzymatic

coo 0

H — C— OH

ch 2 opo 3 ©

3-Phosphoglycerate

Arsenite, (As 0 2 ®) is much more toxic than arsenate. Ar-
senite poisons by an entirely different mechanism than arse-
nate. The arsenic atom of arsenite binds tightly to the two
sulfur atoms of lipoamide (Section 7.12), thereby inhibiting
the enzymes that require this coenzyme.

A Spontaneous hydrolysis of l-arseno-3-phosphoglycerate. Inorganic arsenate

can replace inorganic phosphate as a substrate for glyceraldehyde 3-phosphate A Cary Grant learned about the effects of arsenic in a
dehydrogenase, forming the unstable 1-arseno analog of 1,3-b/sphosphoglycerate. popular 1944 movie.

The formation of ATP by the transfer of a phosphoryl group from a “high energy”
compound (such as 1,3-frisphosphoglycerate) to ADP is termed substrate level phospho-
rylation. This reaction is the first ATP- generating step of glycolysis. It operates at sub-
strate and product concentrations that are close to the equilibrium concentrations. This
is not surprising since the reverse reaction is important in gluconeogenesis, where ATP
is utilized. Flux can proceed easily in either direction.

8. Phosphoglycerate Mutase

Phosphoglycerate mutase catalyzes the near-equilibrium interconversion of 3-phospho-
glycerate and 2-phosphoglycerate.

coo°

H — C — OH

CH 2 0P0 3 © ch 2 oh

3-Phosphoglycerate 2-Phosphoglycerate (1 1 .9)

Mutases are isomerases that catalyze the transfer of a phosphoryl group from one
part of a substrate molecule to another. There are two different types of phosphoglycer-
ate mutase enzymes. In one type, the phosphoryl group is first transferred to an amino

Phosphoglycerate COO^

mutase | ^

< > H — C — OPOo^

1 1 .2 The Ten Steps of Glycolysis

337

acid side chain of the enxyme. The enzyme phosphoryl group is then transferred to the
second site of the substrate molecule. The dephosphorylated intermediate remains
bound in the active site during this process.

Another type of phosphoglycerate mutase makes use of a 2,3-Hsphosphoglycerate
(2,3BPG) intermediate as shown in Figure 11.8. This mechanism also involves a phos-
phorylated enzyme intemediate but it differs from the other type of enzyme because at
no time is there a dephosphorylated metabolite during the reaction. Small amounts of
2,3-frzsphosphoglycerate are required for full activity of this second type of enzyme.
This is because 2,3BPG is required to phosphorylate the enzyme if it becomes dephos-
phorylated. The enzyme will lose its phosphate group whenever 2,3BPG is released
from the active site before it can be converted to 2 -phosphoglycerate or 3 -phosphoglyc-
erate. The second type of phosphoglycerate mutase is called cofactor-dependent PGM,
or dPGM. The first type of enzyme is called cofactor- independent PGM, or iPGM.

dPGM and iPGM are not evolutionarily related. The cofactor-dependent enzyme
(dPGM) belongs to a family of enzymes that include acid phosphatases and fructose
2,6-frzsphosphatase. It is the major form of phosphoglycerate mutase in fungi, some
bacteria, and most animals. The co factor- independent enzyme (iPGM) belongs to the
alkaline phosphatase family of enzymes. This version of phosphoglycerate mutase is
found in plants and some bacteria. Some species of bacteria have both types of enzyme.

▲ Figure 1 1.8

Mechanism of the conversion of 3-phosphoglycerate to 2-phosphoglycerate in animals and fungi. (1) A ly-
sine residue at the active site of phosphoglycerate mutase binds the carboxylate anion of 3-phospho-
glycerate. A histidine residue, which is phosphorylated before the substrate binds, donates its phos-
phoryl group to form the 2,3-b/sphosphoglycerate intermediate. (2) Rephosphorylation of the enzyme
with a phosphoryl group from the C-3 position of the intermediate yields 2-phosphoglycerate.

338

CHAPTER 11 Glycolysis

9. Enolase

2-Phosphoglycerate is dehydrated to phosphoenolpyruvate in a near-equilibrium re-
action catalyzed by enolase. The systematic name of enolase is 2-phosphoglycerate
dehydratase.

coo°

H — C — 0P0 3 ©

Enolase, COO^

Mg© 1

H — C — OH

< - C— OPO 3 © + h

ch 2

2-Phosphoglycerate

Phosphoenolpyruvate

In this reaction, the phosphomonoester 2-phosphoglycerate is converted to an
enol-phosphate ester, phosphoenolpyruvate, by the reversible elimination of water from
C-2 and C-3. Phosphoenolpyruvate has an extremely high phosphoryl group transfer
potential because the phosphoryl group holds pyruvate in its unstable enol form
(Section 10. 7B).

Enolase requires Mg© for activity. Two magnesium ions participate in this reac-
tion: a “conformational” ion binds to the carboxylate group of the substrate, and a
“catalytic” ion participates in the dehydration reaction.

10. Pyruvate Kinase

The second substrate level phosphorylation of glycolysis is catalyzed by pyruvate kinase.
Phosphoryl group transfer to ADP generates ATP in this metabolically irreversible reac-
tion. The unstable enol tautomer of pyruvate is an enzyme-bound intermediate.

COO G

C— 0P0 3 ® +ADP+H® < »

Pyruvate

CH 2 kinase

Phosphoenolpyruvate

coo°

C — OH

L<=h 2 J

Enolpyruvate

coo 0

c = 0 + ATP

ch 3

Pyruvate

( 11 . 11 )

Transfer of the phosphoryl group from phosphoenolpyruvate to ADP is the third
regulated reaction of glycolysis. Pyruvate kinase is regulated both by allosteric modulators
and by covalent modification. In addition, expression of the pyruvate kinase gene in
mammals is regulated by various hormones and nutrients. Recall from Chapter 10 that
phosphoenolpyruvate hydrolysis has a higher standard Gibbs free energy change than
ATP hydrolysis (Table 10.3). Because pyruvate kinase is regulated, the concentration of
phosphoenolpyruvate is maintained at a high enough level to drive ATP formation
during glycolysis.

11.3 The Fate of Pyruvate

The formation of pyruvate from phosphoenolpyruvate is the last step of glycolysis. Further
metabolism of pyruvate typically takes one of five routes (Figure 1 1.9).

1. Pyruvate can be converted to acetyl CoA and acetyl CoA can be used in a number
of metabolic pathways. In one important pathway it is completely oxidized to C0 2
in the citric acid cycle. This fate of pyruvate is described in Chapter 13. This is a
route that operates efficiently in the presence of oxygen.

11.3 The Fate of Pyruvate

339

coo°

c=o

ch 2

coo 0

Oxaloacetate

H 3 N — CH

ch 3

Alanine

Glycolysis

co 2

(2)

CH 3

Pyruvate

(1)

COO

,©

coo 0 ®

^ co 2

S-CoA

c=o

ch 3

Acetyl CoA

HCOH

ch 3

Lactate

CH 2 OH

ch 3

Ethanol

◄ Figure 11.9

Five major fates of pyruvate: (1) Under aerobic
conditions, pyruvate is oxidized to the acetyl
group of acetyl CoA, which can enter the cit-
ric acid cycle for further oxidation. (2) Pyru-
vate can be converted to oxaloacetate, which
can be a precursor in gluconeogenesis.

(3) Under anaerobic conditions, certain
microorganisms ferment glucose to ethanol
via pyruvate. (4) Glucose undergoes anaero-
bic glycolysis to lactate in vigorously exercis-
ing muscles, red blood cells, and certain other
cells. (5) Pyruvate is converted to alanine.

2. Pyruvate can be carboxylated to produce oxaloacetate. Oxaloacetate is one of the citric
acid cycle intermediates but it is also an intermediate in the synthesis of glucose. The
fate of pyruvate as a precursor in gluconeogenesis is covered in Chapter 12.

3. In some species, pyruvate can be reduced to ethanol, which is then excreted from
cells. This reaction normally takes place under anaerobic conditions where entry of
acetyl CoA into the citric acid cycle is unfavorable.

4. In some species, pyruvate can be reduced to lactate. Lactate can be transported to
cells that convert it back to pyruvate for entry into one of the other pathways. This
is also an anaerobic pathway.

5. In all species, pyruvate can be converted to alanine.

During glycolysis, NAD® is reduced to NADH at the glyceraldehyde 3 -phosphate
dehydrogenase reaction (Step 6). In order for glycolysis to operate continuously, the cell
must be able to regenerate NAD®. Otherwise, all the coenzyme would rapidly accumu-
late in the reduced form, and glycolysis would stop. Under aerobic conditions, NADH
can be oxidized by the membrane-associated electron transport system (Chapter 14),
which requires molecular oxygen. Under anaerobic conditions, the synthesis of ethanol
or lactate consumes NADH and regenerates the NAD® essential for continued glycolysis.

The fate of pyruvate as a precursor in
amino acid biosynthesis is discussed
in Chapter 17.

In some species, pyruvate can be
converted to phosphoenolpyruvate
(Section 12.1B).

A. Metabolism of Pyruvate to Ethanol

Many bacteria, and some eukaryotes, are capable of surviving in the absence of oxygen.
They convert pyruvate to a variety of compounds that are secreted. Ethanol is one of
these compounds. It assumes significance in biochemistry because the synthesis of
ethanol by highly selected strains of yeast is important in the production of beer and
wine. Yeast cells convert pyruvate to ethanol and C0 2 and oxidize NADH to NAD®. Two
reactions are required. First, pyruvate is decarboxylated to acetaldehyde in a reaction cat-
alyzed by pyruvate decarboxylase. This enzyme requires the coenzyme thiamine diphos-
phate (TDP); its mechanism was described in the coenzymes chapter (Section 7.7).

Alcohol dehydrogenase catalyzes the reduction of acetaldehyde to ethanol. This
oxidation-reduction reaction is coupled to the oxidation of NADH. These reactions and

KEY CONCEPT

In the absence of oxygen, eukaryotes
have to give up the net gain of 2 NADH
molecules in order to make lactate or
ethanol.

340 CHAPTER 11 Glycolysis

O v

-c

-OH

CH 2 0P0 3 ©

Glyceraldehyde 3-phosphate

-Pi

Glyceraldehyde

3-phosphate

dehydrogenase

NAD® *-

^ NADH + H® ■

OPO,®

H— C— OH

CH 2 OP0 3 ®
1 ,3-B/sphosphoglycerate

coo®

c=o

ch 3

Pyruvate

decarboxylase

H ©

C0 2
O

ch 3

Acetaldehyde

NADH + H®^-

Alcohol

dehydrogenase k NAQ ©

H — C— OH

ch 3

Ethanol

A Figure 11.10

Anaerobic conversion of pyruvate to ethanol
in yeast.

the cycle of NAD® /NADH reduction and oxidation in alcoholic fermentation are shown
in Figure 11.10. Fermentation refers to a process where electrons from glycolysis — in the
form of NADH — are passed to an organic molecule such as ethanol instead of being passed on
to the membrane- associated electron transport chain and ultimately oxygen ( respiration ).
The sum of the glycolytic reactions and the conversion of pyruvate to ethanol is

Glucose + 2 Pi© + 2 ADP© + 2 H® *

2 Ethanol + 2 C0 2 + 2 ATP© + 2 H 2 0 (11.12)

These reactions have familiar commercial roles in the manufacture of beer and bread.
In the brewery, the carbon dioxide produced during the conversion of pyruvate to
ethanol can be captured and used to carbonate the final alcoholic brew; this gas pro-
duces the foamy head. In the bakery, carbon dioxide is the agent that causes bread
dough to rise.

B. Reduction of Pyruvate to Lactate

Pyruvate is reduced to lactate in a reversible reaction catalyzed by lactate dehydro-
genase. This reaction is common in anaerobic bacteria and also in mammals.

coo®

C = O + NADH + H

ch 3

Pyruvate

COO'

Lactate

dehydrogenase

HO — C — H + NAD

CH 3
L-Lactate

(11.13)

Lactate dehydrogenase is a classic dehydrogenase using NAD® as a coenzyme; the
mechanism was presented in Section 7.4. This is an oxidation-reduction reaction in
which pyruvate is reduced to lactate by transfer of a hydride ion from NADH.

The lactate dehydrogenase reaction oxidizes the reducing equivalents generated in
the glyceraldehyde 3 -phosphate reaction and lowers the potential energy gain of glycol-
ysis. It plays the same role that ethanol production accomplishes in other species
(Figure 11.10). The net effect is to maintain flux in the glycolytic pathway and the pro-
duction of ATP. In bacteria, lactate is secreted or converted to other end products, such
as propionate. In mammals, lactate can only be reconverted to pyruvate.

The production of lactate in mammalian cells is essential in tissues where glucose is
the main carbon source and reducing equivalents (NADH) are not needed in biosyn-
thesis reactions or cannot be used to generate ATP by oxidative phosphorylation. A
good example is the formation of lactate in skeletal muscle cells during vigorous exer-
cise. Lactate formed in muscle cells is transported out of cells and carried via the blood-
stream to the liver, where it is converted to pyruvate by the action of hepatic lactate de-
hydrogenase (see Cori cycle, Section 12.2A). Further metabolism of pyruvate requires
oxygen. When the supply of oxygen to tissues is inadequate, all tissues produce lactate
by anaerobic glycolysis.

The overall reaction for glucose degradation to lactate is

Glucose + 2 P|© + 2 ADP© * 2 Lactate© + 2 ATP© + 2 H 2 Q (11.14)

Lactic acid is also produced by Lactobacillus and certain other bacteria when they fer-
ment the sugars in milk. The acid denatures the proteins in milk, causing the curdling
necessary for cheese and yogurt production.

Regardless of the final product — ethanol or lactate — glycolysis generates two mole-
cules of ATP per molecule of glucose consumed. Oxygen is not required in either case.
This feature is essential not only for anaerobic organisms but also for some specialized
cells in multicellular organisms. Some tissues (such as kidney medulla and parts of the
brain), termed obligatory glycolytic tissues, rely on glycolysis for all their energy. In the

11.4 Free Energy Changes in Glycolysis 341

BOX 1 1.4 THE LACTATE OF THE LONG-DISTANCE RUNNER

Most of you have heard stories about lactate buildup during
strenuous exercise. It all sounds so plausible. When muscle
cells are working hard they use up glucose to generate ATP,
which is required for muscle contraction. During very stren-
uous activity, the production of pyruvate may outstrip its
ability to be oxidized by the citric acid cycle. If muscle cells
aren’t getting enough oxygen, then pyruvate is converted to
lactic acid and the accumulation of lactic acid causes acidosis
leading to muscle pain and reduced efficiency.

It’s a nice story, but it’s wrong.

Lactate concentration in muscle cells and in the blood-
stream does increase but lactate is not an acid. It cannot do-
nate a proton, so the increase in protons (acidosis) must
come from another source. Lactate really is the product of
the lactate dehydrogenase reaction, not lactic acid (which can
donate a proton).

There is no net production of protons in the pathway
leading from glucose to lactate. The acidosis seen after stren-
uous exercise is mostly due to the release of protons during
ATP hydrolysis associated with muscle contraction. This is a
temporary imbalance since ATP is soon regenerated in order
to maintain a high steady state concentration. Lactate may
indirectly contribute to some acidosis because, as a potent

anion, it may affect buffering capacity but the effect is not
large. Lactate has been getting a bum rap for decades, includ-
ing previous editions of this textbook.

cornea of the eye, for example, oxygen availability is limited by poor blood circulation.
Anaerobic glycolysis provides the necessary ATP for such tissues.

11.4 Free Energy Changes in Glycolysis

When the glycolytic pathway is operating, the flow of metabolites is from glucose to
pyruvate. Under these conditions, the Gibbs free energy change for every single reaction
must be either negative or zero. It is interesting to compare the standard Gibbs free en-
ergy changes (AG°') and the actual Gibbs free energy changes (AG) under conditions
where flux through the glycolytic pathway is high. Such conditions occur in erythro-
cytes where blood glucose is the main source of energy and there is very little synthesis
of carbohydrates (or any other molecules). The actual concentrations of the intermedi-
ates in glycolysis have been measured and the Gibbs free energy changes have been cal-
culated. The standard Gibbs free energy changes for each of the ten reactions of glycoly-
sis are shown in Table 11.2, The first column lists AG°' values under typical standard
conditions (25°C and zero ionic strength) and the second column corrects those
standard Gibbs free energy changes to mammalian physiological conditions (37°C in the
presence of Mg®, Ca®, Na© and K©).

Figure 11.11 shows the cumulative standard Gibbs free energy changes and actual
free energy changes for the glycolytic reactions in erythrocytes. The vertical axis indicates
cumulative Gibbs free energy changes for each of the steps of glycolysis. The figure illus-
trates the difference between the Gibbs free energy changes under standard physiological
conditions (A G° r ) and actual free energy changes under cellular conditions (AG).

The blue plot tracks the actual cumulative free energy changes. It shows that each
reaction has a Gibbs free energy change that is either negative or zero. This is an essen-
tial requirement for conversion of glucose to pyruvate. It follows that the overall path-
way, which is the sum of the individual reactions, must also have a negative free energy
change. The overall Gibbs free energy change for glycolysis is about —72 kj mol -1 under
the conditions found in erythrocytes.

342

CHAPTER 11 Glycolysis

Table 1 1.2 Standard Gibbs free energies for reactions of glycolysis

Glycolysis

reaction

AG°' (kJ mol 1 )
(standard conditions)

AC°' (kJ mol 1 )
(physiological conditions)

-17.2

-19.4

+2.0

+2.8

-18.0

-15.6

+28.0

+24.6

+7.9

+7.6

+6.7

+2.6

-18.8

-16.4

+4.4

+6.4

-2.7

-4.5

-25.5

-27.2

Data from Minakami and de Verdier (1 976) and Li et al. (201 0).

KEY CONCEPT

The net production of product in a
metabolic pathway (flux) will only occur if:
(a) the overall Gibbs free energy change
is negative, and (b) the Gibbs free energy
change of each step in the pathway is
either negative or zero.

The actual Gibbs free energy changes are large only for Steps 1,3, and 10, which are
catalyzed by hexokinase, phosphofructokinase-1, and pyruvate kinase, respectively —
the steps that are both metabolically irreversible and regulated. The AG values for the
other steps are very close to zero. In other words, these other steps are near- equilibrium
reactions in cells.

In contrast, the standard Gibbs free energy changes for the same ten reactions ex-
hibit no consistent pattern. Although the three reactions with large negative Gibbs free
energy changes in cells also have large standard Gibbs free energy changes, this may be
coincidental since some of the near- equilibrium reactions in cells also have large values
for AG°'. Furthermore, some of the AG°' values for the reactions of glycolysis are posi-
tive, indicating that under standard conditions, flux through these reactions occurs to-
ward substrate rather than product. This is especially obvious in Step 4 (aldolase) and
Step 6 (glycer aldehyde 3 -phosphate dehydrogenase). In other types of cells these near-
equilibrium reactions might operate in the opposite direction during glucose synthesis.

Figure 11.11 ►

Cumulative standard and actual Gibbs free
energy changes for the reactions of glycolysis.

The vertical axis indicates free energy
changes in kJ mol -1 . The reactions of gly-
colysis are plotted in sequence horizontally.
The upper plot (red) tracks the standard free
energy changes, and the bottom plot (blue)
shows actual free energy changes in erythro-
cytes. The interconversion reaction cat-
alyzed by triose phosphate isomerase
(Reaction 5) is not shown. [Adapted from
Hamori, E. (1975). Illustration of free energy
changes in chemical reactions. J. Chem. Ed.
52:370-373.]

Standard Gibbs
free energy changes

Actual Gibbs
> free energy changes
AG about 72 kJ mol 1

11.5 Regulation of Glycolysis

343

11.5 Regulation of Glycolysis

The regulation of glycolysis has been examined more thoroughly than that of any other
pathway. Data on regulation come primarily from two types of biochemical research: en-
zymology and metabolic biochemistry. In enzymological approaches, metabolites are
tested for their effects on isolated enzymes and the structure and regulatory mechanisms
of individual enzymes are studied. Metabolic biochemistry analyzes the concentrations
of pathway intermediates in vivo and stresses pathway dynamics under cellular condi-
tions. We sometimes find that in vitro studies are deceptive as indicators of pathway dy-
namics in vivo. For instance, a compound may modulate enzyme activity in vitro , but
only at concentrations not found in the cell. Accurate interpretation of biochemical data
greatly benefits from a combination of enzymological and metabolic expertise.

In this section, we examine each regulatory site of glycolysis. Our primary focus is
on the regulation of glycolysis in mammalian cells — in particular, those cells where gly-
colysis is an important pathway. Variations on the regulatory themes presented here can
be found in other species.

The regulatory effects of metabolites on glycolysis are summarized in Figure 11.12.
The activation of glycolysis is desirable when ATP is required by processes such as mus-
cle contraction. Hexokinase is inhibited by excess glucose 6-phosphate, and PFK-1 is
inhibited by the accumulation of ATP and citrate (an intermediate in the energy-
producing citric acid cycle). ATP and citrate both signal an adequate energy supply.
Consumption of ATP leads to the accumulation of AMP, which relieves the inhibition
of PFK-1 by ATP. Fructose 2,6-frzsphosphate also relieves this inhibition. The rate of for-
mation of fructose 1,6-frisphosphate then increases, which in certain tissues activates
pyruvate kinase. Glycolytic activity decreases when its products are no longer required.

A. Regulation of Hexose Transporters

The first potential step for regulating glycolysis is the transport of glucose into the cell.
In most mammalian cells, the intracellular glucose concentration is far lower than the
blood glucose concentration, and glucose moves into the cells, down its concentration
gradient, by passive transport. All mammalian cells possess membrane-spanning

Glucose

Hexokinase

N *"- Glucose 6-phosphate

Citrate

ATP ' Fructose 6-phosphate

AMP ► Phosphofructokinase-1

+ t +

Fructose 2,6-b/sphosphate

Fructose 1,6-b/sphosphate

i
I
I
i
I
I

i + +

Pyruvate kinase

Phosphoenolpyruvate

ATP

◄ Figure 11.12

Summary of the metabolic regulation of the
glycolytic pathway in mammals. Not shown
are the effects of ADP on PFK-1, which vary
among species.

Pyruvate

344 CHAPTER 11 Glycolysis

kinase

domains

Insulin

Insulin binds to
cell-surface receptors

Tyrosine-

kinase

domains

Vesicle

▲ Figure 11.13

Regulation of glucose transport by insulin. The

binding of insulin to cell-surface receptors
stimulates intracellular vesicles containing
membrane-embedded GLUT4 transporters
to fuse with the plasma membrane. This de-
livers GLUT4 transporters to the cell surface
and thereby increases the capacity of the
cell to transport glucose.

Membrane transport systems are
described in Section 9.1 1.

glucose transporters. Intestinal and kidney cells have a Na® -dependent cotransport
system called SGLT1 for absorbing dietary glucose and urinary glucose, respectively.
Other mammalian cells contain transporters from the GLUT family of passive hexose
transporters. Each of the six members of the GLUT family has unique properties suit-
able for the metabolic activities of the tissues in which it is found.

The hormone insulin stimulates high rates of glucose uptake into skeletal and heart
muscle cells and adipocytes via the transporter GLUT4. When insulin binds to receptors
on the cell surface, intracellular vesicles that have GLUT4 embedded in their mem-
branes fuse with the cell surface by exocytosis (Section 9.1 ID), thereby increasing the
capacity of the cells to transport glucose (Figure 11.13). Because GLUT4 is found at
high levels only in striated muscle and adipose tissue, insulin- regulated uptake of glu-
cose occurs only in these tissues.

In most tissues, a basal level of glucose transport in the absence of insulin is main-
tained by GLUT1 and GLUT3. GLUT2 transports glucose into and out of the liver,
and GLUT5 transports fructose in the small intestine. GLUT7 transports glucose
6-phosphate from the cytoplasm into the endoplasmic reticulum.

Once inside a cell, glucose is rapidly phosphorylated by the action of hexokinase.
This reaction traps the glucose inside the cell since phosphorylated glucose cannot cross
the plasma membrane. As we will see, phosphorylated glucose can also be used in glyco-
gen synthesis or in the pentose phosphate pathway (Chapter 12).

B. Regulation of Hexokinase

The reaction catalyzed by mammalian hexokinase is metabolically irreversible (because
it is regulated) but in bacteria and many other eukaryotes hexokinase is not regulated.
In those species, the concentrations of reactants and products reach equilibrium. In
mammals, the various forms of hexokinase are subject to complex regulation.

At physiological concentrations, the enzyme product, glucose 6-phosphate,
allosterically inhibits hexokinase isozymes I, II, and III, but not glucokinase (isozyme IV).
Glucokinase is more abundant than the other hexokinases in the liver and the insulin-
secreting cells of the pancreas. The concentration of glucose 6-phosphate increases
when glycolysis is inhibited at sites further along the pathway. The inhibition of hexoki-
nases I, II, and III by glucose 6-phosphate therefore coordinates the activity of hexokinase
with the activity of subsequent enzymes of glycolysis.

Glucokinase is suited to the physiological role of the liver in managing the supply
of glucose for the entire body. In most cells, glucose concentrations are maintained far

11.5 Regulation of Glycolysis 345

BOX 1 1.5 GLUCOSE 6-PHOSPHATE HAS A PIVOTAL METABOLIC ROLE IN THE LIVER

Glucose 6-phosphate is an initial substrate for several metabolic
pathways (figure below). We have already seen that it is the ini-
tial intermediate in glycolysis. Glucose 6-phosphate is formed
rapidly in liver cells from dietary glucose or newly synthesized
glucose (from gluconeogenesis in liver cells; Section 12.1).

The principal use of liver glucose 6-phosphate is to
maintain a constant concentration of blood glucose. Glucose
6 -phosphatase is the enzyme responsible for catalyzing hy-
drolysis of glucose 6-phosphate to glucose. (This reaction is
also the last step in gluconeogenesis.)

Glucose 6-phosphate that is not required for blood glu-
cose is stored as liver glycogen (Section 12.6). Glycogen is

subsequently degraded when a supply of glucose is needed.
Hormones regulate both the synthesis and degradation of
glycogen.

In addition to using it for balancing the blood glucose
concentration, the liver metabolizes glucose 6-phosphate by
the pentose phosphate pathway (Section 12.5) to produce ri-
bose 5-phosphate (for nucleotides) and NADPH (for synthe-
sis of fatty acids). We have seen in this chapter that glucose
6-phosphate can also enter the glycolytic pathway, where it is
converted initially to pyruvate, which leads to another major
metabolite — acetyl CoA.

T Glucose 6-phosphate is at a pivotal position in
carbohydrate metabolism in the liver.

Glucose

-Diet

■i

(Rapid) I Hexokinases

Glucose 6-phosphate

Pentose

Ribose 5-phosphate phosphate
pathway

Glucose

6-phosphatase

NADPH

Glucose for
export to blood

Gluconeogenesis
-Glucose 1-phosphate

k Glycogen

below the concentration in blood. However, glucose freely enters the liver via GLUT2,
and the concentration of glucose in liver cells matches the concentration in blood. The
blood glucose concentration is typically 5 mM, though after a meal it can rise as high as
10 mM. Most hexokinases have K m values for glucose of about 0.1 mM or less. In con-
trast, glucokinase has a K m of 2 to 5 mM for glucose; in addition, it is not significantly
inhibited by glucose 6-phosphate. Therefore, liver cells can form glucose 6-phosphate
(for glycogen synthesis) by the action of glucokinase when glucose is abundant and
other tissues have sufficient glucose.

The activity of glucokinase is modulated by fructose phosphates. In liver cells, a
regulatory protein inhibits glucokinase in the presence of fructose 6-phosphate, lower-
ing its affinity for glucose to about 10 mM (Figure 1 1.14). Note that the v 0 vs. [S] curves
for glucokinase are sigmoidal and not the hyperbolic curves expected for an enzyme
obeying Michaelis-Menten kinetics. This is a common feature of allosterically regulated
proteins. It means that there is no true K m value for glucokinase. We can say that the ef-
fect of the regulatory protein is to raise the apparent K m of the enzyme. Flux through
glucokinase is usually low because liver cells always contain considerable fructose 6-
phosphate. The flux can increase after a meal, when fructose 1 -phosphate — derived
only from dietary fructose — relieves the inhibition of glucokinase by the regulatory
protein. Therefore, the liver can respond to increases in blood carbohydrate concentra-
tions with proportionate increases in the rate of phosphorylation of glucose.

(mM)

C. Regulation of Phosphofructokinase-1

The second site of allosteric regulation is the reaction catalyzed by phosphofructokinase-1.
PFK-1 is a large, oligomeric enzyme with a molecular weight ranging in different
species from about 130,000 to 600,000. The quaternary structure of PFK-1 also varies
among species. The bacterial and mammalian enzymes are both tetramers; the yeast

▲ Figure 11.14

Plot of initial velocity (v 0 ) versus glucose
concentration for glucokinase. The addition of
a regulatory protein lowers the enzyme’s affinity
for glucose. The blood glucose concentration
is 5 to 10 mM.

346 CHAPTER 11 Glycolysis

Figure 1 1.15 ►

Regulation of PFK-1 by ATP and AMP. In the

absence of AMP, PFK-1 is almost completely
inhibited by physiological concentrations of
ATP. In the range of AMP concentrations
found in the cell, the inhibition of PFK-1 by
ATP is almost completely relieved. [Adapted
from Martin, B. R. (1987). Metabolic Regu-
lation: A Molecular Approach (Oxford: Black-
well Scientific Publications), p. 222.]

enzyme is an octamer. This complex enzyme has several regulatory sites. The regulatory
properties of the Escherichia coli phosphofructokinase-1 are described in Section 5.10A.

ATP is both a substrate and, in most species, an allosteric inhibitor of PFK-1. ATP
increases the apparent K m of PFK- 1 for fructose 6-phosphate. The bacterial enzyme is
activated by ADP but in mammals AMP is the allosteric activator of PFK-1. AMP acts
by relieving the inhibition caused by ATP (Figure 11.15). ADP activates mammalian
PFK-1 but inhibits the plant kinase; in bacteria, protists, and fungi, the regulatory ef-
fects of purine nucleotides vary among species.

The concentration of ATP does not change very much in most mammalian cells
despite large changes in the rate of its formation and utilization. However, as discussed in
Section 10.6, significant changes in the concentrations of ADP and AMP do occur because
these molecules are present in cells in much lower concentrations than ATP and small
changes in the level of ATP cause proportionally larger changes in the levels of ADP and
AMR The steady state concentrations of these compounds are therefore able to control
flux through PFK- 1 .

Recall that activation by ADP (or AMP) makes sense in light of the net production
of ATP in glycolysis. Elevated levels of ADP or AMP indicate a deficiency of ATP that
can be offset by increasing the rate of degradation of glucose (Section 5.9A).

Citrate, an intermediate of the citric acid cycle, is another physiologically impor-
tant inhibitor of mammalian PFK-1. An elevated concentration of citrate indicates that
the citric acid cycle is blocked and further production of pyruvate would be pointless.
The regulatory effect of citrate on PFK- 1 is an example of feedback inhibition that reg-
ulates the supply of pyruvate to the citric acid cycle. (Phosphoenolpyruvate, not citrate,
inhibits the bacterial enzyme.)

As shown in Figure 11.12, fructose 2,6-frisphosphate is a potent activator of PFK-1,
effective in the micromolar range. This compound is present in mammals, fungi, and
plants, but not prokaryotes. We will return to the role of fructose 2,6-frisphosphate in
the next chapter after we have described gluconeogenesis and glycogen metabolism.

D. Regulation of Pyruvate Kinase

The third site of allosteric regulation of glycolysis is the reaction catalyzed by pyruvate ki-
nase. Single-cell species, such as bacteria, and protists, have a single pyruvate kinase gene.
The enzyme is allosterically regulated in a simple manner — its activity is affected by pyru-
vate and/or fructose 1,6-Hsphosphate. Regulation is much more complex in mammals
because different organs have different requirements for glucose and glycolysis.

Four different isozymes of pyruvate kinase are present in mammalian tissues. The
isozymes found in liver, kidney, and red blood cells yield a sigmoidal curve when initial
velocity is plotted against phosphoenolpyruvate concentration (Figure 11.16a).
This indicates that PEP is an allosteric activator. These enzymes are also allosterically

1 1 .6 Other Sugars Can Enter Glycolysis 347

(a)

(b)

◄ Figure 11.16

Plots of initial velocity (i/ 0 ) versus phospho-
enolpyruvate concentration for pyruvate
kinase, (a) For isozymes in some cells, the
presence of fructose 1,6-6/sphosphate shifts
the curve to the left, indicating that fructose

1,6-b/sphosphate is an activator of the en-
zymes. (b) When liver or intestinal cells are
incubated with glucagon, pyruvate kinase is
phosphorylated by the action of protein ki-
nase A. The curve shifts to the right, indi-
cating less activity for pyruvate kinase.

activated by fructose 1,6-fcphosphate and inhibited by ATP. In the absence of fructose

1.6- frzsphosphate, physiological concentrations of ATP almost completely inhibit
the isolated enzyme. The presence of fructose 1,6-frzsphosphate — probably the most
important modulator in vivo — shifts the curve to the left. With sufficient fructose

1.6- fcphosphate, the curve becomes hyperbolic. Figure 1 1.16a shows that for a range of
substrate concentrations, enzyme activity is greater in the presence of the allosteric acti-
vator. Recall that fructose 1,6-frisphosphate is the product of the reaction catalyzed by
PFK-1. Its concentration increases when the activity of PFK-1 increases. Since fructose

1.6- frzsphosphate activates pyruvate kinase, the activation of PFK-1 (which catalyzes
Step 3 of the glycolytic pathway) causes subsequent activation of pyruvate kinase (the
last enzyme in the pathway). This is an example of feed-forward activation.

The predominant isozyme of pyruvate kinase found in mammalian liver and intes-
tinal cells is subject to an additional type of regulation, covalent modification by phos-
phorylation. Protein kinase A, which also catalyzes the phosphorylation of PFK-2
(Figure 11.17), catalyzes the phosphorylation of pyruvate kinase. Pyruvate kinase is
less active in the phosphorylated state. The change in kinetic behavior is shown in
Figure 11.16b, which depicts a plot of pyruvate kinase activity in liver and intestinal
cells in the presence and absence of glucagon, a stimulator of protein kinase A. Dephos-
phorylation of pyruvate kinase is catalyzed by a protein phosphatase.

The pyruvate kinase activity of liver cells decreases on starvation and increases
on ingestion of a diet high in carbohydrate. These long term changes are due to
changes in the rate of synthesis of pyruvate kinase and not allosteric regulation or
covalent modification.

A Figure 11.17

Pyruvate kinase from the yeast Saccharomyces
cerevisiae, with the activator fructose
1 ,6-/;/sphosphate (red). The active site is
in the large central domain. [PDB 1A3W]

E. The Pasteur Effect

Louis Pasteur observed that when yeast cells grow anaerobically, they produce much
more ethanol and consume much more glucose than when they grow aerobically. Simi-
larly, skeletal muscle accumulates lactate under anaerobic conditions but not when it
metabolizes glucose aerobically. In both yeast and muscle, the rate of conversion of glu-
cose to pyruvate is much higher under anaerobic conditions. The slowing of glycolysis
in the presence of oxygen is called the Pasteur effect. As we will see in Chapter 13, the
complete aerobic metabolism of a glucose molecule produces much more ATP than the
two molecules of ATP produced by glycolysis alone. Therefore, for any given ATP re-
quirement, fewer glucose molecules must be consumed under aerobic conditions. Cells
sense the state of ATP supply and demand, and they modulate glycolysis by several
mechanisms. For example, the availability of oxygen leads to the inhibition of PFK-1
(and thus glycolysis), probably through an increase in the ATP/AMP ratio.

11.6 Other Sugars Can Enter Glycolysis

Glucose and glucose 6-phosphate are the most common substrates for glycolysis, espe-
cially in vertebrates where glucose is circulated in the bloodstream. However, a variety
of other sugars can be degraded by the glycolytic pathway. In this section, we will see
how sucrose, fructose, lactose, galactose, and mannose can be metabolized.

348 CHAPTER 11 Glycolysis

A Invertase from the yeast Schwanniomyces
occidentalis. The active form of the enzyme
is a dimer of identical subunits. Fructose
(space-filling representation) is bound at the
active site. [PDB 3KF3]

A. Sucrose Is Cleaved to Monosaccharides

The disacharide sucrose can be degraded to its two component monosaccharides: fruc-
tose and glucose. This cleavage is catalyzed by a class of enzymes called sucrases. Invertase
(/3-fructofuranosidease) is one of the most common sucrases. It catalyzes a hydrolytic
cleavage of the glycosidic linkage between the oxygen and the glucose residue, produc-
ing fructose and glucose (Figure 11.18). The glucose residues are then phosphorylated
by hexokinase and the fructose residues enter the pathway as described below.

Some bacteria have a very interesting enzyme called sucrose phosphorylase. It
cleaves sucrose in the presence of inorganic phosphate converting it to a molecule of
fructose and a molecule of glucose 1-phosphate (Figure 11.18). All sugars entering gly-
colysis need to be phosphorylated at some stage and this step almost always involves the
expenditure of one ATP equivalent. Sucrose phosphorylase is an important exception
because it produces glucose 1 -phosphate without spending any ATP currency.

B. Fructose Is Converted to Glyceraldehyde 3-Phosphate

Fructose is phosphorylated to fructose 1 -phosphate by the action of a specific ATP-
dependent fructokinase (Figure 11.19). In mammals, this step occurs in the liver after
fructose has been absorbed in the intestine and transferred in the bloodstream. Fructose
1 -phosphate aldolase catalyzes the cleavage of fructose 1 -phosphate to dihydroxy-
acetone phosphate and glyceraldehyde. The glyceraldehyde is then phosphorylated to
glyceraldehyde 3 -phosphate in a reaction catalyzed by triose kinase, consuming a sec-
ond molecule of ATP. Dihydroxyacetone phosphate is converted to a second molecule of
glyceraldehyde 3 -phosphate by the action of triose phosphate isomerase.

Figure 11.18 ►

Entry of other sugars into glycolysis.

Mannose

ATP -

Lactose

ADP^

Glucose 6-phosphate <e

Galactose
1 -phosphate
uridylyltransferase

ATP

ADP

Phosphoglucomutase

Glucose 1-phosphate

(Sue rase)

Sucrose

Fructose 6-phosphate

Hexokinase

ATP

ADP

ATP -

ADP

phosphorylase]'-* -~ SuC [° se
Fructose ◄ '

Fructokinase

Fructose 1,6-b/sphosphate

/ \

Fructose 1 -phosphate

Dihydroxyacetone phosphate Glyceraldehyde 3-phosphate

1 1 .6 Other Sugars Can Enter Glycolysis 349

ch 2 oh

c = o
I

HO — C — H

H — C— OH

ch 2 oh

Fructose

Fructokinase

ATP ADP

CH 2 0P0 3 ©

C=0

HO — C — H

Fructose

1-phosphate

aldolase

H — C— OH

CH 2 OH

Fructose 1 -phosphate

ch 2 opo 3 ©

c = o

Triose
phosphate
isomerase
>

ch 2 oh

Dihydroxyacetone

phosphate

H O

H — C— OH

ch 2 oh

Triose

kinase

ATP ADP

CH 2 0P0 3 ©
HO — C — H

Glyceraldehyde

3-phosphate

H O

H — C— OH
CH 2 0P0 3 ©

Glyceraldehyde

3-phosphate

The two molecules of glyceraldehyde 3 -phosphate produced can then be metabo-
lized to pyruvate by the remaining steps of glycolysis. The metabolism of one molecule
of fructose to two molecules of pyruvate produces two molecules of ATP and two
molecules of NADH. This is the same yield as the conversion of glucose to pyruvate.
Fructose catabolism bypasses phosphofructokinase- 1 and its associated regulation. Reg-
ulation of pyruvate kinase can still control flux in the pathway.

▲ Figure 11.19

Conversion of fructose to two molecules of
glyceraldehyde 3-phosphate.

C. Galactose Is Converted to Glucose 1 -Phosphate

The disaccharide lactose, present in milk, is a major source of energy for nursing
mammals. In newborns, intestinal lactase catalyzes the hydrolysis of lactose to its com-
ponents, glucose and galactose, both of which are absorbed from the intestine and
transported in the bloodstream.

As shown in Figure 1 1.20, galactose — the C-4 epimer of glucose — can be converted
to glucose 1 -phosphate by a pathway in which the nucleotide sugar UDP-glucose (Sec-
tion 7.2A) is recycled. In the liver, galactokinase catalyzes transfer of a phosphoryl
group from ATP to galactose. The galactose 1 -phosphate formed in this reaction ex-
changes with the glucose 1 -phosphate moiety of UDP-glucose by cleavage of the py-
rophosphate bond of UDP-glucose. This reaction is catalyzed by galactose 1 -phosphate
uridylyltransferase and produces glucose 1 -phosphate and UDP-galactose. Glucose

BOX 1 1.6 A SECRET INGREDIENT

Purified invertase is frequently used in the candy industry to
convert sucrose to fructose and glucose. Fructose is sweeter than
sucrose and therefore more appealing in some food. The liquid,
creamy centers of some chocolates are produced by adding inver-
tase — purified from yeast — to a sucrose mixture. In addition to
tasting sweeter, fructose is much less likely to form crystals. The
catalytic breakdown of sucrose inside the chocolate usually takes
several days or weeks at room temperature.

Look for “invertase” on the labels of food to see more exam-
ples of this industrial application of biochemistry, but keep in
mind that not all liquid centers in chocolates are due to added
invertase.

► Cherry Blossom by Lowney’s (Hershey Canada). The liquid center is due to
the presence of added invertase.

350 CHAPTER 11 Glycolysis

UDP-galactose

A Figure 11.20

Conversion of galactose to glucose
6-phosphate. The metabolic intermediate
UDP-glucose is recycled in the process.

The overall stoichiometry for the pathway is
galactose + ATP -> glucose 6-phosphate
+ ADP.

UDP-Galactose is required for biosyn-
thesis of gangliosides (Section 16.1 1).

1 -phosphate can enter glycolysis after conversion to glucose 6-phosphate in a reaction
catalyzed by phosphoglucomutase. UDP-galactose is recycled to UDP-glucose by the action
of UDP-glucose 4-epimerase.

The conversion of one molecule of galactose to two molecules of pyruvate pro-
duces two molecules of ATP and two molecules of NADH, the same yield as the conver-
sions of glucose and fructose. The required UDP-glucose is formed from glucose and
the ATP equivalent UTP, but only small (catalytic) amounts of it are needed since it is
recycled.

Infants fed an exclusive diet of milk rely on galactose metabolism for about 20% of
their caloric intake. In the most common form of the genetic disorder galactosemia
(the inability to properly metabolize galactose), infants are deficient in galactose
1 -phosphate uridylyltransferase. In such cases, galactose 1 -phosphate accumulates in
the cells and this can lead to a compromise in liver function, recognized by the appearance
of jaundice (yellowing of the skin). The liver damage is potentially fatal. Other effects
include damage to the central nervous system. Screening for galactose 1 -phosphate
uridylyltransferase in the red blood cells of the umbilical cord allows detection of galac-
tosemia at birth. Many of the most severe effects of this genetic deficiency can be mitigated
by a special diet that contains very little galactose and lactose.

The majority of humans undergo a reduction in the level of lactase at about 5 to
7 years of age. This is the normal situation found in most other primates. It parallels the
switch from childhood, where mother’s milk is a major source of nourishment, to
adulthood, where milk is not consumed. In some human populations the production of
lactase is not turned off during adolescence. These populations have acquired a mutant
gene that continues to synthesize lactase in adults. As a result, individuals in these pop-
ulations can consume milk products throughout their lives. Northern European popu-
lations and their descendants have high proportions of lactase-producing adults.

In normal adults, lactose is metabolized by bacteria in the large intestine, with the
production of gases such as C0 2 and H 2 and short-chain acids. The acids can cause diar-
rhea by increasing the ionic strength of the intestinal fluid. Milk and milk products are
usually avoided by people who do not synthesize lactase. Since they do not tolerate diets
rich in milk products, they are said to be lactose intolerant although it’s worth keeping in
mind that this is the normal condition in most mammals, and most humans. Some
lactose- intolerant individuals can eat yogurt, in which the lactose has been partially
hydrolyzed by the action of an endogenous /3-galactosidase of the microorganism in the
yogurt culture. A commercially prepared enzyme supplement that contains /3-galactosidase
from a microorganism can be used to pretreat milk to reduce the lactose content or can
be taken when milk products are ingested by lactase -deficient individuals.

11.7 The Entner-Doudoroff Pathway in Bacteria

351

Mannose

Mannose 6-phosphate

Fructose 6-phosphate

▲ Figure 11.21

Conversion of mannose to fructose 6-phosphate.

D. Mannose Is Converted to Fructose 6-Phosphate

The aldohexose mannose is obtained in the diet from glycoproteins and certain polysac-
charides. Mannose is converted to mannose 6-phosphate by the action of hexokinase. In
order to enter the glycolytic pathway, mannose 6-phosphate undergoes isomerization to
fructose 6-phosphate in a reaction catalyzed by phosphomannose isomerase. These two
reactions are depicted in Figure 11.21.

11.7 The Entner-Doudoroff Pathway
in Bacteria

The classic glycolysis pathway is also called the Embden-Meyerhof-Parnas pathway.
This pathway is found in all eukaryotes and many species of bacteria. However, a large
number of bacterial species do not have phosphofructokinase-1 and cannot convert
glucose 6-phosphate to fructose 1,6-fcphosphate in the hexose stage of glycolysis.

The hexose stage of classic glycolysis can be bypassed by the Entner-Doudoroff
pathway. This pathway begins with the conversion of glucose 6-phosphate to 6-phos-
phogluconate, a reaction that is catalyzed by two enzymes: glucose 6-phosphate dehdro-
genase and 6-phosphogluconolactonase (Figure 11.22). The oxidation of glucose
6-phosphate by glucose 6-phosphate dehydrogenase is coupled to the reduction of
NADP®. The dehydrogenase and 6-phosphogluconolactonase enzymes are common in
almost all species since they are required in the pentose phosphate pathway (Section 12.5).
The Entner-Douderoff pathway is the earliest pathway for glucose degradation. The
classic glycolysis pathway (EMP) evolved later.

6-Phosphogluconate is converted to 2-keto-3-deoxy-6-phosphogluconate (KDPG)
in an unusual dehydration (dehydratase) reaction. KDPG is then split by the action of
KDPG aldolase to one molecule of pyruvate and one molecule of glyceraldehyde
3 -phosphate. Pyruvate is the end product of glycolysis and glyceraldehyde 3 -phosphate
can be converted to another molecule of pyruvate by the triose stage of glycolysis. The
enzymes of the triose stage of the EMP pathway are found in all species since they are
essential for glucose synthesis as well as glycolysis. Note that only one molecule of
glyceraldehyde 3 -phosphate passes down the bottom half of the glycolytic pathway for
every glucose 6-phosphate molecule that enters the Entner-Doudoroff pathway. This
means that only one molecule of ATP is produced for every glucose molecule degraded,
whereas two ATP molecules are synthesized during glycolysis. Two reducing equivalents
(NADH) are produced during glycolysis and two in the ED pathway (NADPH in the
first reaction and one molecule of NADH when glyceraldehyde 3 -phosphate is con-
verted to 1,3-frisphosphoglycerate).

In addition to being the main pathway for glucose degradation in some species, the
Entner-Doudoroff pathway is also important in species that possess a complete Embden-
Meyeroff-Parnas pathway. The Entner-Doudoroff pathway is used in the metabolism of
gluconate and other related organic acids. These metabolites cannot be shunted into the
normal glycolytic pathway. Many bacterial species, including E. coli , can grow on gluconate
as their sole carbon source. Under these conditions the main energy-producing degrada-
tion pathway is the Entner-Doudoroff pathway. The first reaction in the ED pathway pro-
duces NADPH instead of NADH and many species use the glucose 6-phosphate dehydroge-
nase reaction as an important source of NADPH reducing equivalents (Section 12.4).

KEY CONCEPT

The classic glycolysis pathway evolved
millions of years after the Entner-Douderoff
and the gluconeogenesis pathways.

352 CHAPTER 11 Glycolysis

Figure 1 1.22 ►

The Entner-Doudoroff pathway.

In Box 12.2 we discuss metabolic
diseases associated with glucose
6-phosphate dehydrogenase in humans.

Aldolases cleave hexoses to two
3-carbon compounds. KDPG is the
third aldolase we have described.

ch 2 opo 3 ©

Glucose 6-phosphate
dehydrogenase

NADPH NADP©

+ H

6-Phosphogluconolactone
— H 2 0

6-Phosphogluconolactonase

~-*H®

H OH
Glucose 6-phosphate

©r

H — C— OH

OH — C — H

H — C— OH

CH 2 0P0 3 ©

6-Phosphogluconate

H 2 0<

6-Phosphogluconate

dehydratase

ch 2

H — C— OH

ch 2 opo 3 ©

2-Keto-3-deoxy-6-phosphogluconate

(KDPG)

KDPG

Aldolase

'Z/

c=o

ch 3

Pyruvate

'C'

H — C— OH
CH 2 0P0 3 ©

Glyceraldehyde 3-phosphate

Summary

1. Glycolysis is a ten-step pathway in which glucose is catabolized to
pyruvate. Glycolysis can be divided into a hexose stage and a triose
stage. The products of the hexose stage are glyceraldehyde 3 -phos-
phate and dihydroxyacetone phosphate. The triose phosphates inter-
convert, and glyceraldehyde 3-phosphate is metabolized to pyruvate.

2. For each molecule of glucose converted to pyruvate, there is a net
production of two molecules of ATP from ADP + Pj and two mol-
ecules of NAD© are reduced to NADH.

3. Under anaerobic conditions in yeast, pyruvate is metabolized to
ethanol and C0 2 . In some other organisms, pyruvate can be con-
verted to lactate under anaerobic conditions. Both processes use
NADH and regenerate NAD©.

4. The overall Gibbs free energy change for glycolysis is negative.
The steps catalyzed by hexokinase, phosphofructokinase-1, and
pyruvate kinase are metabolically irreversible.

5. Glycolysis is regulated at four steps: the transport of glucose into
some cells and the reactions catalyzed by hexokinase, phospho-
fructokinase-1, and pyruvate kinase.

6. Fructose, galactose, and mannose can enter the glycolytic pathway
via conversion to glycolytic metabolites.

7. The Entner-Doudoroff pathway is an alternate pathway for glu-
cose catabolism in some bacteria.

Problems 353

Problems

1. Calculate the number of ATP molecules obtained from the anaer-
obic conversion of each of the following sugars to lactate: (a)
glucose, (b) fructose, (c) mannose, and (d) sucrose.

2. (a) Show the positions of the six glucose carbons in the two lac-
tate molecules formed by anaerobic glycolysis, (b) Under aerobic
conditions, pyruvate can be decarboxylated to yield acetyl CoA
and C0 2 . Which carbons of glucose must be labeled with 14 C to
yield 14 C0 2 ?

3. If 32 P (i.e., isotopically labeled phosphorus) is added to a cell-
free liver preparation undergoing glycolysis, will this label be di-
rectly incorporated in any glycolytic intermediate or pathway
product?

4 . Huntington’s disease is a member of the “glutamine-repeat”
family of diseases. In middle-aged adults the disease causes neu-
rodegenerative conditions, including involuntary movements and
dementia. The mutated protein (Huntington protein) contains a
polyglutamine region with 40 to 120 glutamines that is thought
to mediate a tight binding of this protein to glyceraldehyde
3 -phosphate dehydrogenase (GAPDH). If the brain relies almost
solely on glucose as an energy source, suggest a role for the Hunt-
ington protein in this disease.

5. Fats (triacylglycerols) are a significant source of stored energy in
animals and are metabolized initially to fatty acids and glycerol.
Glycerol can be phosphorylated by the action of a kinase to pro-
duce glycerol 3 -phosphate, which is oxidized to produce dihy-
droxyacetone phosphate.

(a) Write the reactions for the conversion of glycerol to dihy-
droxyacetone phosphate.

(b) The kinase that acts on the prochiral molecule glycerol is
stereospecific, leading to production of L-glycerol 3-phos-
phate. Which carbons of glycerol 3 -phosphate must be
labeled with 14 C so that aerobic glycolysis yields acetyl CoA
with both carbons labeled?

3 CH 2 OH

Glycerol

6. Tumor cells often lack an extensive capillary network and must
function under conditions of limited oxygen supply. Explain why
these cancer cells take up far more glucose and may overproduce
some glycolytic enzymes.

7. Rapid glycolysis during strenuous exercise provides the ATP
needed for muscle contraction. Since the lactate dehydrogenase
reaction does not produce any ATP, would glycolysis be more effi-
cient if pyruvate rather than lactate were the end product?

8. Why are both hexokinase and phosphofructokinase- 1 inhibited
by an ATP analog in which the oxygen atom joining the f3- and
y-phosphorus atoms is replaced by a methylene group ( — CH 2 — )?

9. The AG°' for the aldolase reaction in muscle is +22.8 kj mol -1 . In
view of this, why does the aldolase reaction proceed in the direc-
tion of glyceraldehyde 3-phosphate and dihydroxyacetone phos-
phate during glycolysis?

10 . For the aldolase reaction, calculate the concentration of fructose
l,6-£hsphosphate if the concentrations of DHAP and G3P were
each: (a) 5 /ulM , (b) 50 ^iM, (c) 500 ^iM.

11. The following plot shows the rate of mammalian phosphofruc-
tokinase- 1 (PFK-1) activity versus fructose 6-phosphate (F6P)
concentration in (a) the presence of ATP, AMP, or both and (b) in
the absence or presence of fructose 2,6-frisphosphate (F26P). Ex-
plain these effects on the reaction rates of PFK- 1 .

[F6P] mM

12 . Draw a diagram showing how increased intracellular [cAMP] af-
fects the activity of pyruvate kinase in mammalian liver cells.

13 . In response to low levels of glucose in the blood, the pancreas
produces glucagon, which triggers the adenylyl cyclase signaling
pathway in liver cells. As a result, flux through the glycolytic path-
way decreases.

(a) Why is it advantageous for glycolysis to decrease in the liver
in response to low blood glucose levels?

(b) How are the effects of glucagon on glycolysis reversed when
the level of glucagon decreases in response to adequate blood
glucose levels?

14 . Chemoautotrophs growing in the ocean will sometimes have all
the enzymes needed for glycolysis even though they will never en-
counter external glucose. Why?

354 CHAPTER 11 Glycolysis

Selected Readings

Metabolism of Glucose

Alberty, R. A. (1996) Recommendations for
nomenclature and tables in biochemical thermo-
dynamics. Eur. J. Biochem. 240:1-14.

Cullis, R M. (1987). Acyl group transfer-phospho-
ryl group transfer. In Enzyme Mechanisms , M. I.
Page and A. Williams, eds. (London: Royal Society
of Chemistry), pp. 178-220.

Hamori, E. (1975). Illustration of free energy
changes in chemical reactions. /. Chem. Ed.
52:370-373.

Hoffmann-Ostenhof, O., ed. (1987). Intermediary
Metabolism (New York: Van Nostrand Reinhold).

Li X, Dash RK, Pradhan RK, Qi F, Thompson M,
Vinnakota KC, Wu F, Yang F, Beard DA. (2010) A
database of thermodynamic quantities for the re-
actions of glycolysis and the tricarboxylic acid
cycle. J Phys Chem B. 1 14:16068-16082.

Minakami S. and de Verdier, C-H. (1976) Colori-
metric study on human erythrocyte glycolysis.

Eur. J. Biochem. 65: 451-460.

Ronimus, R. S., and Morgan, H. W. (2003), Distri-
bution and phylogenies of enzymes of the Embden-
Meyerof-Parnas pathway from archaea and hyper-
thermophilic bacteria support a gluconeogenic
origin of metabolism. Archaea 1:199-221.

Seeholzer, S. H., Jaworowski, A., and Rose, I. A.
(1991). Enolpyruvate: chemical determination as a
pyruvate kinase intermediate. Biochem.
30:727-732.

St- Jean, M., Blonski, C., and Sygush, J. (2009).
Charge stabilization and entropy reduction of
central lysine residues in fructose- Hsphosphate
aldolase. Biochem. 48:4528-453 7.

Regulation of Glycolysis

Depre, C., Rider, M. EL, and Hue, L. (1998). Mech-
anisms of control of heart glycolysis. Eur. J.
Biochem. 258:277-290.

Engstrom, L., Ekman, P., Humble, E., and
Zetterqvist, O. (1987). Pyruvate kinase. In The
Enzymes , Vol. 18, P. D. Boyer and E. Krebs, eds.
(San Diego: Academic Press), pp. 47-75.

Gould, G. W., and Holman, G. D. (1993). The glu-
cose transporter family: structure, function and
tissue-specific expression. Biochem. J. 295:329-341.

Pessin, J. E., Thurmond, D. C., Elmendorf, J. S.,
Coker, K. J., and Okada, S. (1999). Molecular basis
of insulin- stimulated GLUT4 vesicle trafficking.
Location! Location! Location! /. Biol. Chem.
274:2593-2596.

Pilkis, S. J., Claus, T. H., Kurland, I. J., and Lange,
A. J. (1995). 6-Phosphofructo-2-kinase/fructose-
2,6-Hsphosphatase: a metabolic signaling enzyme.
Annu. Rev. Biochem. 64:799-835.

Pilkis, S. J., El-Maghrabi, M. R., and Claus, T. H.
(1988). Hormonal regulation of hepatic gluconeo-
genesis and glycolysis. Annu. Rev. Biochem.
57:755-783.

Pilkis, S. J., and Granner, D. K. (1992). Molecular
physiology of the regulation of hepatic
gluconeogenesis and glycolysis. Annu. Rev. Physiol.
54:885-909.

Van Schaftingen, E. (1993). Glycolysis revisited.
Diabetologia 36:581-588.

Yamada, K., and Noguchi, T. (1999). Nutrient and
hormonal regulation of pyruvate kinase gene
expression. Biochem. J. 337:1-11.

Metabolism of Other Sugars

Alvaro-Benito, M., Polo, A., Gonzalez, B., Fernandez-
Lobato, M., and Sanz-Aparicio, J. (2010). Struc-
tural and kinetic analysis of Schwanniomyces occi-
dentalis invertase reveals a new oligomerization
pattern and the role of its supplementary
domain in substrate binding. /. Biol. Chem.
285:13930-13941; doi:10.1074/jbc.M109.095430

Frey, P. A. (1996). The Leloir pathway: a mechanis-
tic imperative for three enzymes to change the
stereochemical configuration of a single carbon in
galactose. FASEB J. 10:461-470.

Itan, Y., Jones, B. L., Ingram, C. J. E., Swallow, D. M.,
and Thomas, M. G. (2010). A worldwide correla-
tion of lactase persistence phenotypes and geno-
types. BMC Evol. Biol. 10:36; www.biomedcentral.
com/1471-2148/10/36

o c

_ o

° o o o

° o

o o

° c

o o

Gluconeogenesis,

The Pentose Phosphate Pathway,
and Glycogen Metabolism

W e have seen that the catabolism of glucose is central to energy metabolism in
some cells. In contrast, all species can synthesize glucose from simple two-
carbon and three-carbon precursors by gluconeogenesis (literally, the forma-
tion of new glucose). Some species, notably photosynthetic organisms, can make these
precursors by fixing carbon dioxide leading to the net synthesis of glucose from inorganic
compounds. In our discussion of gluconeogenesis in this chapter we must keep in mind
that every glucose molecule used in glycolysis had to be synthesized in some species.

The pathway for gluconeogenesis shares some steps with glycolysis, the pathway for
glucose degradation, but four reactions specific to the gluconeogenic pathway are not
found in the degradation pathway. These reactions replace the metabolically irreversible
reactions of glycolysis. These opposing sets of reactions are an example of separate, reg-
ulated pathways for synthesis and degradation (Section 10.2).

In addition to fueling the production of ATP (via glycolysis and the citric acid
cycle), glucose is also a precursor of the ribose and deoxyribose moieties of nucleotides
and deoxynucleo tides. The pentose phosphate pathway is responsible for the synthesis
of ribose as well as the production of reducing equivalents in the form of NADPH.

Glucose availability is controlled by regulating the uptake and synthesis of glucose
and related molecules and by regulating the synthesis and degradation of storage poly-
saccharides composed of glucose residues. Glucose is stored as glycogen in bacteria and
animals and as starch in plants. Glycogen and starch can be degraded to release glucose
monomers that can fuel energy production via glycolysis or serve as precursors in
biosynthesis reactions. The metabolism of glycogen will illustrate another example of
opposing, regulated pathways.

Although the reaction we had found
would be viewed today as utterly
trivial it came nevertheless as a
great surprise ; because , at that time ;
nobody could imagine that the phos-
phorylation of an enzyme could be
involved in its regulation.

— Eddy Fischer, Memories of
Ed Krebs (201 0)

Top: The Cori ester, a-D-glucopyranose 1-phosphate.

355

356

CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

In the next section, we discuss how
other precursors enter the pathway.

In mammals, gluconeogenesis, the pentose phosphate pathway, and glycogen me-
tabolism are closely and coordinately regulated in accordance with the moment-
to- moment requirements of the organism. In this chapter, we review these pathways and
examine some of the mechanisms for regulating glucose metabolism in mammalian cells.
The regulation of glucose and glycogen metabolism in mammals is important from a his-
torical perspective because it was the first example of a signal transduction mechanism.

12.1 Gluconeogenesis

As stated in the introduction, all organisms have a pathway for glucose biosynthesis, or
gluconeogenesis. This is true even for animals that use exogenous glucose as an impor-
tant energy source because glucose may not always be available from external sources or
intracellular stores. For example, large mammals that have not eaten for 16 to 24 hours
have depleted their liver glycogen reserves and need to synthesize glucose to stay alive
because glucose is required for the metabolism of certain tissues, for example, brain.
Some mammalian tissues, primarily liver and kidney, can synthesize glucose from simple
precursors such as lactate and alanine. Under fasting conditions, gluconeogenesis sup-
plies almost all of the body’s glucose. When exercising under anaerobic conditions,
muscle converts glucose to pyruvate and lactate, which travel to the liver and are con-
verted to glucose. Brain and muscle consume much of the newly formed glucose. Bacte-
ria can convert many nutrients to phosphate esters of glucose and to glycogen.

It is convenient to consider pyruvate as the starting point for the synthesis of glu-
cose. The pathway for gluconeogenesis from pyruvate is compared to the glycolytic
pathway in Figure 12.1. Note that many of the intermediates and enzymes are identi-
cal. All seven of the near-equilibrium reactions of glycolysis proceed in the reverse
direction during gluconeogenesis. Enzymatic reactions unique to gluconeogenesis are
required for the three metabolically irreversible reactions of glycolysis. These irre-
versible glycolytic reactions are catalyzed by pyruvate kinase, phosphofructokinase-1,
and hexokinase. In the biosynthesis direction these reactions are catalyzed by different
enzymes.

Although all species have a gluconeogenesis pathway, they don’t all have the glycol-
ysis pathway (Section 1 1.7). This is especially true of bacterial species that diverged very
early in the evolution of prokaryotes. Thus, it seems like gluconeogenesis is the more
ancient pathway, which makes sense since there has to be a source of glucose before
pathways for its degradation can evolve. Since the biosynthesis pathway evolved first, it
is appropriate to think of the glycolytic enzymes as bypass enzymes. These enzymes, es-
pecially phosphofructokinase-1, evolved in order to bypass the metabolically irre-
versible reactions of gluconeogenesis.

The synthesis of one molecule of glucose from two molecules of pyruvate requires
four ATP and two GTP molecules as well as two molecules of NADH. The net equation
for gluconeogenesis is

2 Pyruvate + 2 NADH + 4 ATP + 2 GTP + 6 H 2 0 + 2 H© >

Glucose + 2 NAD© + 4 ADP + 2 GDP + 6 P, (12.1)

Four ATP equivalents are needed to overcome the thermodynamic barrier to the forma-
tion of two molecules of the energy- rich compound phosphoenolpyruvate from two
molecules of pyruvate. Recall that in glycolysis the conversion of phosphoenolpyruvate
to pyruvate is a metabolically irreversible reaction catalyzed by pyruvate kinase. In the
catabolic direction this reaction is coupled to the synthesis of ATP. Two ATP molecules
are required to carry out the reverse of the glycolytic reaction catalyzed by phosphoglyc-
erate kinase. In the hexose stage of gluconeogenesis, no energy is recovered in the steps
that convert fructose 1,6-fcphosphate to glucose because fructose 1,6-Hsphosphate is
not a “high energy” intermediate. Recall that glycolysis consumes two ATP molecules and
generates four, for a net yield of two ATP equivalents and two molecules of NADH. Con-
trast this with the synthesis of one molecule of glucose by gluconeogenesis consuming a
total of six ATP equivalents and two molecules of NADH. As expected, the biosynthesis
of glucose requires energy and its degradation releases energy.

12.1 Gluconeogenesis 357

Glucose

6-phosphatase

Glucose

Glycolysis

ATP

Hexokinase

ADP

Glucose 6-phosphate

Fructose 6-phosphate

ATP

Fructose

1,6-b/sphosphatase

Phosphofructokinase-1

ADP

Fructose 1,6-b/sphosphate

◄ Figure 12.1

Comparison of gluconeogenesis and glycolysis.

There are four metabolically irreversible
reactions of gluconeogenesis (blue). These
are the reactions catalyzed by three different
enzymes in glycolysis (red). Both pathways
include a triose stage and a hexose stage.
Two molecules of pyruvate are therefore
required to produce one molecule of
glucose.

Dihydroxyacetone

phosphate

Glyceraldehyde

3-phosphate

NAD® + ^
NADH + H©

NAD© + P,
NADH + H©

1 ,3-£/sphosphoglycerate

ADP

ATP

3-Phosphoglycerate

2-Phosphoglycerate

Pyruvate carboxylase is a biotin-
containing enzyme. The reaction mech-
anism was described in Section 7.10.

Phosphoenolpyruvate
(ADP) GDP^

Phosphoenolpyruvate

carboxykinase /

(ATP) GTP — ^7

Oxaloacetate
ADP + Pi

Pyruvate
carboxylase

Gluconeogenesis ATP

Pyruvate

ADP

Pyruvate

kinase

ATP

A. Pyruvate Carboxylase

We begin our examination of the individual steps in the conversion of pyruvate to glu-
cose with the two enzymes required for synthesis of phosphoenolpyruvate. The two
steps involve a carboxylation followed by decarboxylation. In the first step, pyruvate
carboxylase catalyzes the conversion of pyruvate to oxaloacetate. The reaction is cou-
pled to the hydrolysis of one molecule of ATP (Figure 12.2).

Pyruvate carboxylase is a large, complex, enzyme composed of four identical sub-
units. Each subunit has a biotin prosthetic group covalently linked to a lysine residue.
The biotin is required for the addition of bicarbonate to pyruvate. Pyruvate carboxylase
catalyzes a metabolically irreversible reaction — it can be allosterically activated by acetyl
CoA. This is the only regulatory mechanism known for the enzyme. Accumulation of

coo°

1 ©

C = 0 + ATP + HC0 3 u

£. H Bicarbonate
Pyruvate

Pyruvate

carboxylase

coo©

C = 0 + ADP + P|

ch 2

coo

Oxaloacetate

▲ Figure 12.2

Pyruvate carboxylase reaction.

358 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

coo 0

c=o

ch 2

coo°

Oxaloacetate

GTP

(ATP)

Phosphoenolpyruvate
carboxykinase (PEPCK)

(12.3)

COO 0

C — 0P0 3 ® + GDP(ADP)

II + co 2

ch 2

Phosphoenolpyruvate

(PEP)

▲ Figure 12.3

Phosphoenolpyruvate carboxykinase reaction.

▲ Phosphoenolpyruvate carboxykinase from
rat ( Rattus norvegicus). The closed active site
contains a bound GTP molecule, a molecule
of oxaloacetate, and two Mn© ions (pink).
[PDB 3DT4]

acetyl CoA indicates that it is not being efficiently metabolized by the citric acid cycle.
Under these conditions, pyruvate carboxylase is stimulated in order to direct pyruvate
to oxaloacetate instead of acetyl CoA. Oxaloacetate can enter the citric acid cycle or
serve as a precursor for glucose biosynthesis.

Bicarbonate is one of the substrates in the reaction shown in Figure 12.2. Bicarbonate
is formed when carbon dioxide dissolves in water so the reaction is sometimes written
with C0 2 as a substrate. The pyruvate carboxylase reaction plays an important role in
fixing carbon dioxide in bacteria and some eukaryotes. This role may not be so obvious
when we examine gluconeogenesis since the carbon dioxide is released in the very next
reaction; however, much of the oxaloacetate that is made is not used for gluconeogenesis.
Instead, it replenishes the pool of citric acid cycle intermediates that serve as precursors
to the biosynthesis of amino acids and lipids (Section 13.7).

B. Phosphoenolpyruvate Carboxykinase

Phosphoenolpyruvate carboxykinase (PEPCK) catalyzes the conversion of oxaloacetate
to phosphoenolpyruvate (Figure 12.3). This is a well-studied enzyme with an induced-
fit binding mechanism similar to that described for yeast hexokinase (Section 6.5C) and
citrate synthase (Section 13. 3 A).

There are two different versions of PEPCK. The enzyme found in bacteria, protists,
fungi, and plants uses ATP as the phosphoryl group donor in the decarboxylation reac-
tion. The animal version uses GTP. In most species, the enzyme displays no allosteric
kinetic properties and has no known physiological modulators. Its activity is most often
affected by controls at the level of transcription of its gene. The level of PEPCK activity in
cells influences the rate of gluconeogenesis. This is especially true in mammals where glu-
coneogenesis is mostly confined to cells in the liver, kidneys, and small intestine. During
fasting in mammals, prolonged release of glucagon from the pancreas leads to continued
elevation of intracellular cAMP, that triggers increased transcription of the PEPCK gene
in the liver and increased synthesis of PEPCK. After several hours, the amount of PEPCK
rises and the rate of gluconeogenesis increases. Insulin, abundant in the fed state, acts in
opposition to glucagon at the level of the gene reducing the rate of synthesis of PEPCK.

The two-step synthesis of phosphoenolpyruvate from pyruvate is common in most
eukaryotes, including humans. This is the main reason why it’s usually shown when the
gluconeogenesis pathway is described (Figure 12.1). However, many species of bacteria
can convert pyruvate directly to phosphoenolpyruvate in an ATP-dependent reaction cat-
alyzed by phosphoenolpyruvate synthetase (Figure 12.4). The products of this reaction
include AMP and Pi. The second phosphoryl from ATP is transferred to pyruvate. Thus,
two ATP equivalents are used in the conversion of pyruvate to phosphoenolpyruvate.
This is a much more efficient route than the eukaryotic two-step pathway catalyzed by
pyruvate carboxylase and PEPCK. The presence of phosphoenolpyruvate synthetase in
bacterial cells is due to the fact that efficient gluconeogenesis is much more important
in bacteria than in eukaryotes.

C. Fructose 1 ,6-6/sphosphatase

The reactions of gluconeogenesis between phosphoenolpyruvate and fructose 1,6-
frisphosphate are simply the reverse of the near- equilibrium reactions of glycolysis. The
next reaction in the glycolysis pathway — catalyzed by phosphofructokinase-1 — is meta-
bolically irreversible. In the biosynthesis direction, this reaction is catalyzed by the third
enzyme specific to gluconeogenesis, fructose 1,6-frisphosphatase. This enzyme catalyzes
the conversion of fructose 1,6-frisphosphate to fructose 6-phosphate.

( 12 . 2 )

12.1 Gluconeogenesis 359

BOX 12.1 SUPERMOUSE

Richard Hansons group at Case Western Reserve University in
Cleveland, Ohio, USA, created a form of supermouse by adding
extra copies of the cytoplasmic phosphoenopyruvate carboxy-
kinase gene. The homozygous transgenic mice expressed 10X
more PEPCK in their skeletal muscle. They were hyperactive,
aggressive, and capable of running for extended periods of time
on a mouse treadmill (up to 5 km without stopping!). They ate
more than control mice but were significantly smaller.

The rodent athletes converted prodigious amounts of
oxaloacetate into phosphoenolpyruvate and subsequently to
intermediates in the gluconeogenesis pathway, including glu-
cose. Their muscle cells had many more mitochondria than
the cells of normal mice.

The biochemical explanation of this hyperactivity is not
completely understood. Its probably due to effects on the citric
acid cycle (Chapter 13). This allows increased flux in that
pathway leading ultimately to higher levels of ATP. When
asked whether this genetic modification would be a good way

of creating superior human athletes, Hanson and Hakimi
(2008) replied, “The PEKCK-C mus mice are very aggressive;
the world needs less , not more aggression,” besides, the cre-
ation of such transgenic humans is “. . . neither ethical nor
possible.”

Watch the video at: youtube.com/watch?v=4PXC_mctsgY

▲ Mighty Mouse © CBS Operations.

As you might expect, hydrolysis of the phosphate ester in this reaction is associated with
a large negative standard Gibbs free energy change ( AG°'). The actual Gibbs free energy
change in vivo is also negative because this reaction is metabolically irreversible. The
mammalian enzyme displays sigmoidal kinetics and is allosterically inhibited by AMP
and by the regulatory molecule fructose 2,6-frisphosphate. Thus, the reaction cannot
reach equilibrium. Recall that fructose 2,6-frisphosphate is a potent activator of phos-
phofructokinase-1, the enzyme that catalyzes the formation of fructose 1,6-frisphosphate
in glycolysis (Section 1 1.5C). The two enzymes that catalyze the interconversion of fructose
6-phosphate and fructose 1,6-Hsphosphate are reciprocally controlled by the concen-
tration of fructose 2,6-frisphosphate (see Section 12.6C).

D. Glucose 6-phosphatase

The final step of gluconeogenesis is the hydrolysis of glucose 6-phosphate to form glu-
cose. The enzyme is glucose 6-phosphatase.

v Figure 12.4

Phosphoenolpyruvate synthetase reaction.

coo 0

C=0 + ATP

ch 3

Pyruvate

Phosphoenolpyruvate

synthetase

coo 0

C — 0P0 3 © + ATP + AMP

II + P;

ch 2

Phosphoenolpyruvate

(PEP)

Additional effects of glucagon and
insulin are described in Section 12.6C.

Although we present glucose as the final product of gluconeogenesis, this is not true
in all species. In most cases, the biosynthetic pathway ends with glucose 6-phosphate.
This product is an activated form of glucose. It becomes the substrate for additional
carbohydrate pathways leading to synthesis of glycogen (Section 12.6), starch and su-
crose (Section 15.11), pentose sugars (Section 12.5), and other hexoses.

In mammals, glucose is an important end product of gluconeogenesis since it
serves as an energy source for glycolysis in many tissues. Glucose is made in the cells of
the liver, kidneys, and small intestine and exported to the bloodstream. In these cells,
glucose 6-phosphatase is bound to the endoplasmic reticulum with its active site in the
lumen. The enzyme is part of a complex that includes a glucose 6-phosphate trans-
porter (G6PT) and a phosphate transporter. G6PT moves glucose 6-phosphate from the

360 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Defects in the activities of glucose
6-phosphatase or glucose 6-phosphate
transporter cause von Gierke disease
(Section 12.8).

cytosol to the interior of the ER where it is hydrolyzed to glucose and inorganic phos-
phate. Phosphate is returned to the cytosol and glucose is transported to the cell surface
(and the bloodstream) via the secretory pathway.

The other enzymes required for gluconeogenesis are found, at least in small
amounts, in many mammalian tissues. Glucose 6-phosphatase is found only in cells from
the liver, kidneys, and small intestine, so only these tissues can synthesize free glucose.
Cells of tissues that lack glucose 6-phosphatase retain glucose 6-phosphate for internal
carbohydrate metabolism.

KEY CONCEPT

Mammalian fuel metabolism is an
important subset of biochemistry because
it helps us to understand our own bodies.

12.2 Precursors for Gluconeogenesis

The main substrates for glucose 6 -phosphate synthesis are pyruvate, citric acid cycle inter-
mediates, three-carbon intermediates in the pathway (e.g. glyceraldehyde 3-phosphate),
and two -carbon compounds such as acetyl Co A. Acetyl Co A is converted to oxaloacetate in
the glyoxylate cycle, that operates in bacteria, protists, fungi, plants, and some animals
(Section 13.8). Some organisms can fix inorganic carbon by incorporating it into two-
carbon and three-carbon organic compounds (e.g., Calvin cycle, Section 15.4). These com-
pounds enter the gluconeogenesis pathway resulting in net synthesis of glucose from C0 2 .

Mammalian biochemistry is focused on fuel metabolism and biosynthesis of glu-
cose from simple precursors and is it usually discussed in that context. The major gluco-
neogenic precursors in mammals are lactate and most amino acids, especially alanine.
Glycerol, which is produced from the hydrolysis of triacylglycerols, is also a substrate for
gluconeogenesis. Glycerol enters the pathway after conversion to dihydroxyacetone
phosphate. Precursors arising in nongluconeogenic tissues must first be transported to
the liver to be substrates for gluconeogenesis.

A. Lactate

Glycolysis generates large amounts of lactate in active muscle and red blood cells. Lactate
from these and other sources enters the bloodstream and travels to the liver where it is
converted to pyruvate by the action of lactate dehydrogenase. Pyruvate can then be a sub-
strate for gluconeogenesis. Glucose produced by the liver enters the bloodstream for deliv-
ery to peripheral tissues, including muscle and red blood cells. This sequence is known as
the Cori cycle (Figure 12.5). The conversion of lactate to glucose requires energy, most of
which is derived from the oxidation of fatty acids in the liver. Thus, the Cori cycle transfers
chemical potential energy in the form of glucose from the liver to the peripheral tissues.

B. Amino Acids

The carbon skeletons of most amino acids are catabolized to pyruvate or intermediates of
the citric acid cycle. The end products of these catabolic pathways can serve directly as pre-
cursors for synthesis of glucose 6-phosphate in cells that are capable of gluconeogenesis. In
peripheral mammalian tissues, pyruvate formed from glycolysis or amino acid catabolism

Figure 12.5 ►

Cori cycle. Glucose is converted to L-lactate
in muscle cells. Some of this lactate is se-
creted and passes via the bloodstream to
the liver. Lactate is converted to glucose in
the liver and the glucose is secreted into the
bloodstream where it is taken up by muscle
cells. Both tissues are capable of synthesiz-
ing glycogen and mobilizing it.

LIVER MUSCLE

12.2 Precursors for Gluconeogenesis 361

must be transported to the liver before it can be used in glucose synthesis. The Cori cycle is
one way of accomplishing this transfer by converting pyruvate to lactate in muscle and re-
converting it to pyruvate in liver cells. The glucose-alanine cycle is a similar transport system
(Section 17.9B). Pyruvate can also accept an amino group from an a-amino acid, such as
glutamate, forming alanine by the process of transamination (Section 7.2B) (Figure 12.6).

Alanine travels to the liver, where it undergoes transamination with a-ketoglutarate
to re-form pyruvate for gluconeogenesis. Amino acids become a major source of carbon
for gluconeogenesis during fasting when glycogen supplies are depleted.

The carbon skeleton of aspartate is also a precursor of glucose. Aspartate is the
amino group donor in the urea cycle, a pathway that eliminates excess nitrogen from
the cell (Section 17.9B). Aspartate is converted to fumarate in the urea cycle and then
fumarate is hydrated to malate that is oxidized to oxaloacetate. In addition, the
transamination of aspartate with a-ketoglutarate directly generates oxaloacetate.

C. Glycerol

The catabolism of triacylglycerols produces glycerol and acetyl CoA. As mentioned earlier,
acetyl CoA contributes to the net formation of glucose through reactions of the glyoxy-
late cycle (Section 13.8). The glyoxylate cycle does not contribute to net synthesis of
glucose from lipids in mammalian cells. Glycerol, however, can be converted to glucose
by a route that begins with phosphorylation to glycerol 3 -phosphate, catalyzed by glyc-
erol kinase (Figure 12.7). Glycerol 3 -phosphate enters gluconeogenesis after conversion
to dihydroxyacetone phosphate. This oxidation can be catalyzed by a flavin containing
glycerol 3 -phosphate dehydrogenase complex embedded in the inner mitochondrial
membrane. The outer face of this enzyme binds glycerol 3 -phosphate and electrons are
passed to ubiquinone (Q) and subsequently to the rest of the membrane-associated
electron transport chain. The oxidation of glycerol 3 -phosphate can also be catalyzed by
the NAD® requiring cytosolic glycerol 3 -phosphate dehydrogenase, although this en-
zyme is usually associated with the reverse reaction for making glycerol. Both enzymes
are found in the liver, the site of most gluconeogenesis in mammals.

D. Propionate and Lactate

In ruminants — cattle, sheep, giraffes, deer, and camels — the propionate and lactate pro-
duced by the microorganisms in the rumen (chambered stomach) are absorbed and

Glucose

t
t

Gluconeogenesis

Glycerol

ATP

ADP

NADH,H©

z' — > Dihydroxyacetone
f phosphate

Glycerol

kinase

Cytosolic
glycerol
k 3-phosphate

NAD® deh V dro 9 enase

Glycerol 3-phosphate

CYTOSOL

INNER

MITOCHONDRIAL

MEMBRANE

MITOCHONDRIAL MATRIX

▲ Figure 12.7

Gluconeogenesis from glycerol. Glycerol 3-phosphate can be oxidized by a glycerol 3-phosphate de-
hydrogenase complex in the mitochondrial membrane. A cytoplasmic version of this enzyme inter-
converts dihydroxyacetone phosphate and glycerol 3-phosphate.

COO'

COO 0 H 3 N — C — H

C = 0 +

ch 3

Pyruvate

CH 2

Glutamate

Transamination

coo 0

-u

H 3 N — CH +

ch 2

ch 3

oh 2

Alanine

S \ n

0 0°

u-Ketoglutarate

a Figure 12.6

Conversion of pyruvate to alanine. Pyruvate
can be converted to alanine in peripheral
tissues. Alanine is secreted into the blood-
stream where it is taken up by liver cells
and converted back to pyruvate by the same
transamination reaction. Pyruvate then
serves as a precursor for gluconeogenesis.

▲ Glycerol 3-phosphate dehydrogenase. This
is the human ( Homo sapiens ) version of the
cytosolic enzyme containing DHAP and
NAD© at the active site. The structure of
the membrane-bound version is not known.
[PDB 1WPQ]

362 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

► Precursors for gluconeogenesis. The gly-
oxylate pathway, the Calvin cycle, and fixation
of CO 2 into acetate, do not occur in mammals.
Propionate is produced by microorganisms
in the rumen of ruminants.

Triacylglycerols

Glycerol

Amino acids

Propionate

Glucose

Glucose 6-phosphate

C0 2

Calvin

Glyceraldehyde 3-phosphate c y cle
Dihydroxyacetone phosphate

Gluconeogenesis

Phosphoenolpyruvate

Oxaloacetate

Pyruvate

Amino acids

Lactate

Acetyl CoA

Fatty acids Acetate <— C0 2

enter the gluconeogenesis pathway. Propionate is converted to propionyl CoA and then
to succinyl CoA. These reactions will be covered in the chapter on lipid metabolism
(Section 16.3). Succinyl CoA is an intermediate of the citric acid cycle that can be me-
tabolized to oxaloacetate. Lactate from the rumen is oxidized to pyruvate.

E. Acetate

Many species can utilize acetate as their main source of carbon. They can convert
acetate to acetyl CoA that serves as the precursor to oxaloacetate. Bacteria and

BOX 12.2 GLUCOSE IS SOMETIMES CONVERTED TO SORBITOL

In most animals, glucose — whether from gluconeogenesis,
food, or glycogenolysis — is usually oxidized or reincorpo-
rated into glycogen. However, in some mammalian tissues
(including, testes, pancreas, brain and the lens of the eye),
glucose can be converted to fructose as shown in the pathway
below. Aldose reductase catalyzes the reduction of glucose to
produce sorbitol and polyol dehydrogenase catalyzes the oxi-
dation of sorbitol to fructose. This short pathway supplies es-
sential fructose for some cells. For example, fructose is the
main fuel for sperm cells.

Aldose reductase has a high K m value for glucose so flux
through this pathway is normally low and glucose is usually
metabolized by glycolysis. When the concentration of glucose is
higher than usual (e.g., in individuals with diabetes), increased
amounts of sorbitol are produced in tissues such as the lens.
There is less polyol dehydrogenase activity than aldose reduc-
tase activity so sorbitol can accumulate. Since membranes are
relatively impermeable to sorbitol, the resulting change in the
osmolarity of the cells causes aggregation and precipitation of
lens proteins leading to cataracts — opaque regions in the lens.

t Production of sorbitol from glucose.

H v°

ch 2 oh

H — C — OH

H — C— OH

c=o

HO — C— H

Aldose

reductase

HO — C — H

Polyol

dehydrogenase

HO — C — H

H — C — OH

NADPH + H© J

H— C— OH

NAD©^

H— C— OH

— u-
1

NADP©

H — C— OH

NADH + H©

H — C — OH

CH 2 OH

Glucose

Sorbitol

Fructose

12.3 Regulation of Gluconeogenesis 363

Glucose

AMP

Citrate ATP

Fructose

1,6-jb/'sphosphatase

Fructose

1,6-b/sphosphate

Fructose 2,6-
b/sphosphate

Gluconeogenesis

Glycolysis

Phosphofractokinase-1

+ t + t

i i

AMP

Fructose 2,6-
b/sphosphate

◄ Figure 12.8

Regulation of glycolysis and gluconeogenesis
by metabolites. The interconversions of fruc-
tose 6-phosphate/fructose 1,6-b/sphosphate
and phosphoenol pyruvate/pyruvate are cat-
alyzed by different metabolically irreversible
enzymes. Changing the activity of any of the
enzymes can affect not only the rate of flux
but also the direction of flux toward either
glycolysis or gluconeogenesis. The net effect
is enhanced regulation at the expense of the
hydrolysis of ATP.

Phosphoenol

pep pyruvate

carboxykinase / r 3

Oxaloacetate

Pyruvate

kinase

F--

Acetyl CoA

Pyruvate ^

J q

- ATP

- Phosphorylation catalyzed
by protein kinase A

Fructose 1,6-b/sphosphate

Lactate

single-celled eukaryotes such as yeast utilize acetate as a precursor for gluconeogene-
sis. Some species of bacteria can synthesize acetate directly from C0 2 . In those species
the gluconeogenesis pathway provides a route for the synthesis of glucose from inor-
ganic substrates.

12.3 Regulation of Gluconeogenesis

Gluconeogenesis is carefully regulated in vivo. Glycolysis and gluconeogenesis are opposing
catabolic and anabolic pathways that share some enzymatic steps but certain reactions are
unique to each pathway. For example, phosphofructokinase-1 catalyzes a reaction in glycol-
ysis and fructose 1,6-fcphosphatase catalyzes the opposing reaction in gluconeogenesis;
both reactions are metabolically irreversible. Usually, only one of the enzymes is active at
any given time.

Short-term regulation of gluconeogenesis (regulation that occurs within min-
utes and does not involve the synthesis of new protein) is exerted at two sites — the
reactions involving pyruvate and phosphoenolpyruvate and those that intercon-
vert fructose 1,6-frisphosphate and fructose 6-phosphate (Figure 12.8). When there
are two enzymes catalyzing the same reaction (in different directions), modulating
the activity of either enzyme can alter the flux through the two opposing pathways.
For example, inhibiting phosphofructokinase-1 stimulates gluconeogenesis since
more fructose 6-phosphate enters the pathway leading to glucose rather than being
converted to fructose 1,6-^zsphosphate. Simultaneous control of fructose 1,6-
frzsphosphatase also regulates the flux of fructose 1,6-^zsphosphate toward either
glycolysis or gluconeogenesis.

We’ve encountered phosphofructokinase-1 (PFK-1) several times, most notably in
the previous chapter (Section 11. 5C) and in our discussion of allostery (Section 5.9).
Now it’s time to examine the effect of the allosteric effector, fructose 2,6-frzsphosphate,
on the activity of PFK-1.

Fructose 2,6-Hsphosphate is formed from fructose 6-phosphate by the action of the
enzyme phosphofructokinase-2 (PFK-2) (Figure 12.9). In mammalian liver, a different

/3-D-Fructose 6-phosphate

/3-D-Fructose 2,6-b/sphosphate
▲ Figure 12.9

Interconversion of j8-D-fructose 6-phosphate
and /?-D-fructose 2,6-b/sphosphate.

364 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

BOX 12.3 THE EVOLUTION OF A COMPLEX ENZYME

Bacterial versions of phosphofructokinase-1 are homote-
tramers (Figure 5.19). The functional unit is a head-to-tail
dimer with two active sites and two regulatory sites in the in-
terface between the monomers. Phosphoenolpyruvate (PEP)
inhibits the enzyme.

In eukaryotes, a tandem gene duplication occurred in
the fungi/animal lineage. This was followed by a fusion of the
two genes leading to a monomer that was twice the size of the
bacterial version. This larger monomer resembled the bacter-
ial dimer with two active sites and two regulatory sites. Over a

period of millions of years these sites became modified. One
of the active sites continued to bind fructose 6-phosphate and
ATP catalyzing the formation of fructose 1,6-frzsphosphate. In
the reverse reaction it binds fructose 1,6-frzsphosphate. The
other active site evolved to bind fructose 2,6-frzsphosphate,
which became an allosteric activator.

The two original regulatory sites also evolved to accom-
modate new ligands. Citrate became the new inhibitor at one
of the sites and the other site became the allosteric site for
regulation by ATP (inhibitor) or AMP (activator).

▼ Evolution of the fungal and animal versions of phosphofructokinase-1.

Bacterial

enzyme

PEP

Active

site

Active

site

Active

site

Eukaryotic

enzyme

Citrate

ATP

AMP

Fructose 2,6-b/sphosphate

▲ T conformation (inactive) of fructose

1 .6- Zz/sphosphatase. This is the tetrameric
enzyme from human (Homo sapiens) bound
to the allosteric inhibitor AMP (space-filling)
at the regulatory sites between the two
dimers. The competitive inhibitor fructose

2.6- b/'sphosphate (bal l-and-stick) is bound at
the active sites of each monomer. [PDB 1EYJ]

active site on the same protein catalyzes the hydrolytic dephosphorylation of fructose

2.6- frisphosphate, re-forming fructose 6-phosphate. This activity of the enzyme is called
fructose 2,6-frisphosphatase. The dual activities of this bifunctional enzyme control the
steady state concentration of fructose 2,6-frzsphosphate and, ultimately, the switch
between glycolysis and gluconeogenesis.

As shown in Figure 12.8, the allosteric effector fructose 2,6-frisphosphate acti-
vates PFK-1 and inhibits fructose 1,6-frzsphosphatase. Note that an increase in fructose

2.6- Hsphosphate has reciprocal effects: it stimulates glycolysis and inhibits gluconeogen-
esis. Similarly, AMP affects the two enzymes in a reciprocal manner; inhibiting fructose

1.6- frisphosphatase and activating phosphofructokinase-1. The regulation of the bifunc-
tional enzyme PFK-2/fructose 2,6-frzsphosphatase will be described after we cover
glycogen metabolism.

12.4 The Pentose Phosphate Pathway

The pentose phosphate pathway is a pathway for the synthesis of three pentose phosphates:
ribulose 5 -phosphate, ribose 5 -phosphate, and xylulose 5 -phosphate. Ribose 5 -phosphate
is required for the synthesis of RNA and DNA. The complete pathway has two stages: an
oxidative stage and a nonoxidative stage (Figure 12.10). In the oxidative stage, NADPH
is produced when glucose 6-phosphate is converted to the five-carbon compound ribulose
5-phosphate.

Glucose 6-phosphate + 2 NADP© + H 2 0 >

Ribulose 5-phosphate + 2 NADPH + C0 2 + 2 H© (12.4)

12.4 The Pentose Phosphate Pathway

365

(a)

Glucose 6-phosphate

Glucose

6-phosphate

dehydrogenase

NADP®

^ NADPH + H®

6-Phosphogluconolactone

^-h 2 o

6-Phosphogluconolactonase

H ©

6-Phosphogluconate

NADP®

6-Phosphogluconate v A1Ar ^ nil
dehydrogenase ^ NADPH

^ C0 2

Ribulose 5-phosphate

Ribulose

5-phosphate

3-epimerase

„ J

Ribose

5-phosphate

isomerase

V ^

Oxidative

stage

(b) 6C (3)

1C (3)

5C (3)

5C (2) 5C (1)

Xylulose Ribose

5-phosphate 5-phosphate

Glyceraldehyde Fructose Fructose

3-phosphate 6-phosphate 6-phosphate

Non-

oxidative

stage

▲ Figure 12.10

Pentose phosphate pathway, (a) The oxidative
stage of the pathway produces a five-carbon
sugar phosphate, ribulose 5-phosphate,
with concomitant production of NADPH.

The nonoxidative stage produces the
glycolytic intermediates glyceraldehyde
3-phosphate and fructose 6-phosphate.

(b) The path of carbon in the pentose phos-
phate pathway. In the oxidative stage, three
molecules of a six-carbon compound are
converted to three molecules of a five-
carbon sugar (ribulose 5-phosphate) with
release of three molecules of CO 2 . In the
nonoxidative stage, three molecules of five-
carbon sugars are interconverted to pro-
duce two molecules of a six-carbon sugar
(fructose 6-phosphate) and one molecule of
a three-carbon compound (glyceraldehyde
3-phosphate).

If a cell requires both NADPH and nucleotides then all the ribulose 5 -phosphate is
isomerized to ribose 5 -phosphate and the pathway is completed at this stage. In some
cases, more NADPH than ribose 5-phosphate is needed and most of the pentose phos-
phates are converted to intermediates in the gluconeogenesis pathway.

The nonoxidative stage of the pentose phosphate pathway disposes of the pen-
tose phosphate formed in the oxidative stage by providing a route to gluconeogenesis

366 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Glucose 6-phosphate

Glucose

6-phosphate

dehydrogenase

NADP®

^ NADPH + H®

6-Phosphogluconolactone

6-Phospho-

gluconolactonase

^-h 2 0

H ©

®o o

H— C— OH

HO — C — H

H— C— OH

CH 2 0P0 3 ©

6-Phosphogluconate

dehydrogenase

NADP®
^ NADPH
^C0 2

CH 2 OH

c = o
I

H— C— OH

ch 2 opo 3 ©

Ribulose 5-phosphate
▲ Figure 12.11

Oxidative stage of the pentose phosphate path-
way. Two molecules of NADP© are reduced
to two molecules of NADPH for each mole-
cule of glucose 6-phosphate that enters the
pathway.

or glycolysis. In this stage, ribulose 5-phosphate is converted to the intermediates
fructose 6-phosphate and glyceraldehyde 3 -phosphate. If all the pentose phosphate
were converted to these intermediates, the sum of the nonoxidative reactions would
be the conversion of three pentose molecules to two hexose molecules plus one
triose molecule.

3 Ribulose 5-phosphate >

2 Fructose 6-phosphate + Glyceraldehyde 3-phosphate (12.5)

Both fructose 6-phosphate and glyceraldehyde 3-phosphate can be metabolized by glycolysis
or gluconeogenesis.

Lets take a closer look at the individual reactions of the pentose phosphate pathway.

A. Oxidative Stage

The three reactions of the oxidative stage of the pentose phosphate pathway are shown in
Figure 12.1 1. The first two steps are the same as those in the bacterial Entner-Doudoroff
pathway (Section 1 1.7). The first reaction, catalyzed by glucose 6-phosphate dehydroge-
nase (G6PDH), is the oxidation of glucose 6-phosphate to 6-phosphogluconolactone.
This step is the major regulatory site for the entire pentose phosphate pathway. Glucose
6-phosphate dehydrogenase is allosterically inhibited by NADPH (feedback inhibition).
This simple regulatory feature ensures that the production of NADPH by the pentose
phosphate pathway is self-limiting.

The next enzyme of the oxidative phase is 6-phosphogluconolactonase that cat-
alyzes the hydrolysis of 6-phosphogluconolactone to the sugar acid 6-phosphogluconate.
Finally, 6-phosphogluconate dehydrogenase catalyzes the oxidative decarboxylation of
6-phosphogluconate. This reaction produces a second molecule of NADPH, ribulose
5 -phosphate, and C0 2 . In the oxidative stage, therefore, a six- carbon sugar is oxidized
to a five-carbon sugar plus C0 2 and two molecules of NADP® are reduced to two mol-
ecules of NADPH.

B. Nonoxidative Stage

The nonoxidative stage of the pentose phosphate pathway consists entirely of near equilib-
rium reactions. This stage of the pathway provides five-carbon sugars for biosynthesis and
introduces sugar phosphates into glycolysis or gluconeogenesis. Ribulose 5-phosphate has
two fates: an epimerase can catalyze the formation of xylulose 5-phosphate, or an iso-
merase can catalyze the formation of ribose 5-phosphate (Figure 12.12). (Note the differ-
ence between an epimerase and an isomerase.) Ribose 5-phosphate is the precursor of
the ribose (or deoxyribose) portion of nucleotides. The remaining steps of the pathway
convert the five- carbon sugars into glycolytic intermediates. Rapidly dividing cells that
require both ribose 5-phosphate (as a precursor of ribonucleotide and deoxyribonu-
cleotide residues) and NADPH (for the reduction of ribonucleotides to deoxyribonu-
cleotides) generally have high pentose phosphate pathway activity.

The overall pentose phosphate pathway (Figure 12.10) shows that in the nonoxida-
tive stage two molecules of xylulose 5 -phosphate and one molecule of ribose 5 -phosphate
are interconverted to generate one three-carbon molecule (glyceraldehyde 3 -phosphate)
and two six-carbon molecules (fructose 6-phosphate). Thus, the carbon-containing
products from the passage of three molecules of glucose through the pentose phosphate
pathway are glyceraldehyde 3-phosphate, fructose 6-phosphate, and C0 2 . The balanced
equation for this process is

3 Glucose 6-phosphate + 6 NADP® + 3 H 2 0 > 2 Fructose 6-phosphate +

Glyceraldehyde 3-phosphate + 6 NADPH + 3 GQ 2 + 6 H® (12.6)

12.4 The Pentose Phosphate Pathway 367

BOX 12.4 GLUCOSE 6-PHOSPHATE DEHYDROGENASE DEFICIENCY IN HUMANS

The genetics of human glucose 6 -phosphate dehydrogenase has
been the subject of much research. There are two different en-
zymes that can catalyze the reaction shown in Figure 12.1 1. One
of the genes (G6PDH) is found on the X chromosome (Xq28)
and it is expressed almost exclusively in red blood cells. The
other gene (H6PDH) encodes an enzyme that is less specific; it
can use other hexose substrates. Hexose 6-phosphate dehydro-
genase is synthesized in many cells where it serves as the first en-
zyme in the oxidative stage of the pentose phosphate pathway.

The glucose 6-phosphate dehydrogenase reaction is the
only reaction capable of reducing NADP© in red blood cells;
consequently, deficiencies of this enzyme have drastic effects
on the metabolism of these cells. Other cells are not affected
since they contain H6PDH. G6PDH deficiency in humans
causes hemolytic anemia.

There are hundreds of different alleles of the X chromo-
some G6PDH gene. The variants produce lower amounts of
the enzyme or they alter its catalytic efficiency. There are no
known null mutants in the human population because the
complete absence of G6PDH activity is lethal. Note that
males are more likely to be affected since they have only a
single copy of the gene on their one X chromosome.

It is estimated that 400 million people have some form of
G6PDH deficiency and suffer from mild forms of hemolytic
anemia. The symptoms can be life threatening if the patient is
treated with certain drugs that are normally prescribed for
other diseases. Many of these individuals have an increased re-
sistance to malaria because the malarial parasite does not sur-
vive well in red blood cells that produce lowered amounts of
NADPH. This explains why there are so many deficiency alleles

segregating in the human population in spite of the fact that
the pentose phosphate pathway is inefficient. Its an example of
balanced selection like the familiar sickle cell anemia example.

Human genome database entries for these genes can be
viewed on the Entrez Gene website [ncbi.nlm.nih.gov/gene].
Type in the entries for the G6PDH gene (2531) or the H6PDH
gene (9563). The Online Mendelian Inheritance in Man (OMIM)
webpage is at ncbi.nlm.nih.gov/omim. The entry for G6PDH is
MIM=305900 and the entry for H6PDH is MIM= 138090.

▲ Human glucose 6-phosphate dehydrogenase, variant Canton R459L. The

enzyme is a dimer of dimers (tetramer). Two molecules of NADP© are
bound at the active sites in each dimer. [PDB 1QK1]

ch 2 oh
c = o

ch 2 oh

CH 2 OH

c— o e

C = 0

c— OH

< » HO— C — H

Ribulose

5-phosphate

H — C— OH

H— C — OH

3-epimerase

1 ^

CH 2 0P0 3 cy

ch 2 opo 3 (

2,3-Enediol

Xylulose

intermediate

5-phosphate

H— C — OH

ch 2 opo 3 ®

Ribose

Ribulose

5-phosphate

isomerase

H OH

11 ©

C— 0°

H — C— OH

ch 2 opo 3 ®

1,2-Enediol

intermediate

H .0

H — C — OH

CH 2 0P0 3 ®

Ribose

5-phosphate

The reactions of the nonoxidative stage
of the pentose phosphate pathway are
similar to those of the regeneration stage
of the reductive pentose phosphate cycle
of photosynthesis (Section 15.8).

◄ Figure 12.12

Conversion of ribulose 5-phosphate to xylulose
5-phosphate or ribose 5-phosphate. In either
case, the removal of a proton forms an ene-
diol intermediate. Reprotonation forms ei-
ther the ketose xylulose 5-phosphate or the
aldose ribose 5-phosphate.

368 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

▲ Transketolase from Escherichia coli.

The active site of each monomer contains
one molecule of xylulose 5-phosphate
(space-filling) and the TDP cofactor. [PDN
2R80]

Figure 12.13 y

Reaction catalyzed by transketolase. The

reversible transfer of a glycoaldehyde group
(shown in red) from xylulose 5-phosphate
to ribose 5-phosphate generates glyceralde-
hyde 3-phosphate and sedoheptulose 7-
phosphate. Note that the ketose-phosphate
substrate (in either direction) is shortened
by two carbon atoms, whereas the
aldose-phosphate substrate is lengthened
by two carbon atoms. In this example,

In most cells, the glyceraldehyde 3 -phosphate and fructose 6-phosphate produced
by the pentose phosphate pathway are used to resynthesize glucose 6-phosphate. This
glucose 6 -phosphate molecule can reenter the pentose phosphate pathway. In that case,
the equivalent of one molecule of glucose is completely oxidized to C0 2 by six passages
through the pathway. After six molecules of glucose 6-phosphate are oxidized, the six
ribulose 5-phosphates produced can be rearranged by the reactions of the pentose phos-
phate pathway and part of the gluconeogenic pathway to form five glucose 6-phosphate
molecules. (Recall that two glyceraldehyde 3 -phosphate molecules are equivalent to one
fructose 1,6-frzsphosphate molecule.) If we disregard H 2 0 and H®, the overall stoi-
chiometry for this process is

6 Glucose 6-phosphate + 12 NADP® >

5 Glucose 6-phosphate + 12 NADPH + 6 C0 2 + Pj (12.7)

This net reaction emphasizes that most of the glucose 6-phosphate entering the pentose
phosphate pathway could be recycled; one -sixth is converted to C0 2 and Pi. Indeed, an
alternate name for the pathway is the pentose phosphate cycle.

C. Interconversions Catalyzed by Transketolase
and Transaldolase

The interconversions of the nonoxidative stage of the pentose phosphate pathway are
catalyzed by two enymes called transketolase and transaldolase. These enzymes have
broad substrate specificities.

Transketolase is also called glycoaldehydetransferase. It is a thiamine diphosphate
(TDP) -dependent enzyme that catalyzes the transfer of a two-carbon glycoaldehyde
group from a ketose phosphate to an aldose phosphate. The ketose phosphate is
shortened by two carbons and the aldose phosphate is lengthened by two carbons
(Figure 12.13).

Transaldolase is also called dihydroxyacetonetransferase. It catalyzes the transfer of
a three-carbon dihydroxyacetone group from a ketose phosphate to an aldose phosphate.
The transaldolase reaction of the pentose phosphate pathway converts sedoheptulose
7-phosphate and glyceraldehyde 3 -phosphate to erythrose 4-phosphate and fructose
6-phosphate (Figure 12.14).

5C + 5C

3C + 7C.

ch 2 oh

0 H

CH 2 OH

C = 0
1

HO— C — H

c=o

H — C — OH

0 K H

H — C — OH

)— c — H +

H — r — OH « 7

H — C — OH

1 — C— OH

Transketolase

H — C — OH

H — C— OH

H — C — OH

ch 2 opo 3 ©

ch 2 opo 3 (?

Xylulose

Ribose

Glyceraldehyde

Sedoheptulose

5-phosphate

3-phosphate

7-phosphate

12.5 Glycogen Metabolism

Glucose is stored as the intracellular polysaccharides starch and glycogen. In Chapter 15
we discuss starch metabolism, which occurs mostly in plants. Glycogen is an important
storage polysaccharide in bacteria, protists, fungi and animals. Large glycogen particles
can be easily seen in the cytoplasm of these organisms. Most of the glycogen in verte-
brates is found in muscle and liver cells. Muscle glycogen appears in electron micrographs
as cytosolic granules with a diameter of 10 to 40 nm, about the size of ribosomes.
Glycogen particles in liver cells are about three times larger. The glycogen particles in
bacteria are smaller.

12.5 Glycogen Metabolism

369

ch 2 oh

c=o

HO — C — H

0 H

CH 2 OH
C = 0

H — C— OH

H O

HO — C — H

H — C— OH +

H — C— OH

H — C— OH +

H — C— OH

H — C — OH

Transaldolase

H— C— OH

ch 2 opo 3 ©

CH 2 0P0 3 ©

CH 2 OPO:

Sedoheptulose

Glyceraldehyde

Erythrose

Fructose

7-phosphate

3-phosphate

4-phosphate

6-phosphate

▲ Figure 12.14

Reaction catalyzed by transaldolase. The reversible transfer of a three-carbon dihydroxyacetone group
(shown in red) from sedoheptulose 7-phosphate, to C-l of glyceraldehyde 3-phosphate generates a
new ketose phosphate, fructose 6-phosphate, and releases a new aldose phosphate, erythrose 4-
phosphate. Note that the carbon atoms balance: 7C + 3C 6C + 4C.

A. Glycogen Synthesis

De novo glycogen synthesis requires a preexisting primer of four to eight a-( 1 —> 4)-
linked glucose residues. This primer is attached to a specific tyrosine residue of a pro-
tein called glycogenin (Figure 12.15) via the 1 -hydroxyl group of the reducing end of
the short polysaccharide. The primer is formed in two steps. The first glucose residue is
attached to glycogenin by the action of a glucosyltransferase activity that requires UDP-
glucose. Glycogenin itself catalyzes this reaction as well as the extension of the primer
by up to seven more glucose residues. Thus, glycogenin is both a protein scaffold for
glycogen and an enzyme. Each glycogen molecule (which can contain thousands of glu-
cose residues) contains a single glycogenin protein at its center.

Further glycogen addition reactions begin with glucose 6-phosphate that can be
converted to glucose 1 -phosphate. We saw in Section 11.5 that glucose 6-phosphate
can enter a number of pathways, including glycolysis and the pentose phosphate path-
way. Glycogen synthesis and degradation is mostly a way of storing glucose 6-phos-
phate until it is needed by the cell. The synthesis and degradation of glycogen require
separate enzymatic steps. We have already noted that it is a general rule of metabolism
that biosynthesis pathways and degradation pathways follow different routes.

Three separate enzyme-catalyzed reactions are required to incorporate a mole-
cule of glucose 6-phosphate into glycogen (Figure 12.16). First, phosphoglucomutase
catalyzes the conversion of glucose 6-phosphate to glucose 1 -phosphate. Glucose
1 -phosphate is then activated by reaction with UTP, forming UDP-glucose and py-
rophosphate (PPi). In the third step, glycogen synthase catalyzes the addition of glucose
residues from UDP-glucose to the nonreducing end of glycogen.

Phosphoglucomutase is a ubiquitous enzyme. It catalyzes a near- equilibrium reac-
tion that converts a-D-glucose 6-phosphate to a-D-glucose 1 -phosphate Glucose 1-
phosphate is the famous “Cori ester” discovered by Gerty Cori and Carl Cori in the
1930s when the reactions of glycogen metabolism were first being elucidated.

( 12 . 12 )

u-D-Glucose 6-phosphate

u-D-Glucose 1 -phosphate

▲ Gerty Cori, (1896-1957) biochemist. Carl
Cori and Gerty Cori won the Nobel Prize in
1947 “for their discovery of the course of the
catalytic conversion of glycogen.” This stamp
depicts the “Cori ester” but it’s slightly dif-
ferent than the structure we usually see in
textbooks. Can you spot the difference?

▲ Figure 12.15

Glycogenin from rabbit ( Oryctolagus cuniculus).

The molecule is a homodimer and each of
the active sites contains a bound molecule
of UDP-glucose. [PDB 1LL2]

370 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

▲ Large glycogen particles in a section of a
liver cell. (Electron micrograph.)

▲ Stained glycogen granules in bacteria
( Candidates spp.)

Glucose 6-phosphate

Phosphoglucomutase

Glucose 1 -phosphate

UDP-glucose
pyrophosphorylase

UDP-glucose

__ Glycogen
Glycogen ' (n residues)
synthase

^UDP

Glycogen
(n + 1 residues)

▲ Figure 12.16

Synthesis of glycogen in eukaryotes.

UTP

PP, -> 2 Pi

The mechanism of this reaction is similar to that of cofactor- dependent phospho-
glycerate mutase (Section 11.2 8). Glucose 6-phosphate binds to the phosphoenzyme,
and glucose 1,6-frisphosphate is formed as an enzyme-bound intermediate. Transfer of
the C-6 phosphate to the enzyme leaves glucose 1 -phosphate.

Glucose 1 -phosphate is activated by formation of UDP-glucose in the second step of
glycogen synthesis. In this reaction a UMP group from UTP is transferred to the phos-
phate at C-l with release of pyrophosphate (see Figure 7.6). The enzyme that catalyzes
this reaction is called UDP glucose pyrophosphorylase and it is present in most eukaryotic
species. Note that the activation of glucose requires UTP. The
energy is stored in UDP-glucose where it can be used in many biosynthesis reactions. We
saw in Section 1 1.6 that UDP-glucose can be a substrate for synthesis of UDP-galactose.
(UDP-galactose is used in the synthesis of gangliosides.) The standard Gibbs free energy
change in the UDP glucose pyrophospholylase reaction is close to zero. Under the steady
state, near-equilibrium conditions found in vivo , AG = 0 and the concentrations of glu-
cose 1 -phosphate and UDP-glucose are nearly equal. Flux in the direction of UDP-glucose
synthesis is driven by subsequent hydrolysis of pyrophosphate (Section 10.6). Two ATP
equivalents (UTP and PPj) are used in the activation of glucose.

Glycogen synthesis is a polymerization reaction where glucose units are added one
at a time to a growing polysaccharide chain. This reaction is catalyzed by glucogen syn-
thase (Figure 12.17). Many polymerization reactions are processive — the enzyme
remains bound to the end of the growing chain and addition reactions are very rapid
(see Section 20. 2B). The glycogen synthase reaction is distributive — the enzyme releases
the growing glycogen chain after each reaction.

Glycogen synthases that use UDP-glucose as their substrate are present in protists,
animals, and fungi. Some bacteria synthesize glycogen using ADP-glucose. Starch syn-
thesis in plants also requires ADP-glucose. The glycogen synthase reaction is the major
regulatory step of glycogen synthesis. In animals, there are hormones that control the
rate of glycogen synthesis by altering the activity of glycogen synthase. We will describe
regulation in the next section.

Another enzyme, amylo-(l,4 — > l,6)-transglycosylase, catalyzes branch formation in
glycogen. This enzyme, also known as the branching enzyme, removes an oligosaccharide of
at least six residues from the nonreducing end of an elongated chain and attaches it by an
a-( l— >6) linkage to a position at least four glucose residues from the nearest a-(l — » 6)
branch point. These branches provide many sites for adding or removing glucose residues,
thereby contributing to the speed with which glycogen can be synthesized or degraded.

The complete glycogen molecule has many layers of polysaccharide chains extend-
ing out from the glycogenin core (Figure 12.18). The large granules in liver cells, for ex-
ample, have glycogen molecules with up to 120,000 glucose residues. There are usually
two branches per chain and each chain is 8-14 residues in length. The molecule has
about 12 layers of chains. If there were on average two branches per chain then each
polysacharide unit would have thousands of free ends.

B. Glycogen Degradation

The glucose residues of starch and glycogen are released from storage polymers through
the action of enzymes called polysaccharide phosphorylases: starch phosphorylase (in
plants) and glycogen phosphorylase (in other organisms). These enzymes catalyze the
removal of glucose residues from the nonreducing ends of starch or glycogen, provided
the monomers are attached by a-(l — > 4) linkages. As the name implies, the enzymes
catalyze phosphorolysis — cleavage of a bond by group transfer to an oxygen atom of
phosphate. In contrast to hydrolysis (group transfer to water), phosphorolysis pro-
duces phosphate esters. Thus, the first product of polysaccharide breakdown is a-D-glucose
1 -phosphate (the Cori ester), not free glucose.

Polysaccharide
Polysaccharide + phosphorylase

(n residues) 1

Polysaccharide

, ' . , x + Glucose 1 -phosphate

(n - 1 residues)

(12.9)

12.5 Glycogen Metabolism

371

o 0 o 0

UDP-glucose

Glycogen synthase

Glycogen (n residues)

0 ,

O — P — O— P — O — Uridine
O 0 O 0

UDP

CH 2 OH

CFLOH

▲ Figure 12.17

The glycogen synthase reaction.

The phosphorolysis reaction catalyzed by glycogen phosphorylase is shown in
Figure 12.19. Pyridoxal phosphate (PLP) is a prosthetic group in the active site of
the enzyme. The phosphate group of PLP appears to relay a proton to the substrate
phosphate to help cleave the scissile C — O bond of glycogen. Note that glycogen phospho-
rylase catalyzes a remarkable reaction since it only uses glycogen and inorganic phosphates
as substrates in a reaction that produces a relatively “high energy” compound, glucose
1 -phosphate (Table 10.1).

Glycogen phosphorylase is a dimer of identical subunits. The catalytic sites lie in
the middle of each subunit. It binds phosphate and the end of a glycogen chain
(Figure 12.20). The large glycogen particle binds to a nearby site and the chain being
degraded passes along a groove on the surface of the enzyme. Four or five glucose
residues can be cleaved sequentially before the enzyme has to release a glycogen particle
and re-bind. Thus, in contrast to glycogen synthase, glycogen phosphorylase is partially
processive.

The enzyme stops four glucose residues from a branch point (an a-( 1 —>6) glucosidic
bond) leaving a limit dextrin. The limit dextrin can be further degraded by the action
of the bifunctional glycogen debranching enzyme (Figure 12.21). A glucano transferase
activity of the debranching enzyme catalyzes the relocation of a chain of three glucose
residues from a branch to a free 4-hydroxyl end of the glycogen molecule. Both the orig-
inal linkage and the new one are a- (l —> 4). The other activity of glycogen debranching
enzyme, amylo-l,6-glucosidase, catalyzes hydrolytic (not phosphorolytic) removal of
the remaining a-{ \ —> 6) -linked glucose residue. The products are one free glucose
molecule and an elongated chain that is again a substrate for glycogen phosphorylase.
When a glucose molecule released from glycogen by the action of the debranching en-
zyme enters glycolysis, two ATP molecules are produced (Section 11.1). In contrast,
each glucose molecule mobilized by the action of glycogen phosphorylase (representing
about 90% of the residues in glycogen) yields three ATP molecules. The energy yield
from glycogen is higher than from free glucose because glycogen phosphorylase cat-
alyzes phosphorolysis rather than hydrolysis — no ATP is consumed as in the hexokinase-
catalyzed phosphorylation of free glucose.

The product of glycogen degradation, glucose 1 -phosphate, is rapidly converted to
glucose 6-phosphate by phosphoglucomutase.

▲ Figure 12.18

A glycogen molecule. Two polysaccharides
(blue) are attached to each core glycogenin
molecule. Each chain core has 8-14
residues and two branches. Not all branches
are shown. Seven layers are numbered but
typical glycogen molecules have 8-12 lay-
ers, depending on the species.

There’s no magical net gain of energy
by storing glucose as glycogen since
the cost of incorporating glucose
6-phosphate into glycogen is two ATP
equivalents (Figure 12.16).

372 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

▲ Inhibiting glycogen phosphorylase.

The action of glycogen phosphorylase pro-
duces glucose in the liver. Insulin controls
this activity by inactivating glycogen phos-
phorylase but in the absence of insulin (e.g.,
Type II diabetes), excess production of glu-
cose can be dangerous. Many inhibitors of
glycogen phosphorylase have been devel-
oped as possible treatments for diabetes.
One of them is a cyclic maltose molecule
shown here bound to the active sites of the
rabbit ( Oryctolagus cuniculis ) enzyme.
[PDB1P2G]

Allosteric

site

Glycogen-
binding site

▲ Figure 12.20

Binding and catalytic sites on glycogen
phosphorylase.

Glycogen (n residues)

°0— P — O— H

Glycogen

phosphorylase

CFLOH CH 2 OH

▲ Figure 12.19

Cleavage of a glucose residue from the nonreducing end of a glycogen chain, catalyzed by glycogen
phosphorylase.

12.6 Regulation of Glycogen Metabolism in Mammals

Mammalian glycogen stores glucose in times of plenty (after feeding, a time of high glu-
cose levels) and supplies glucose in times of need (during fasting or in “fight or flight”
situations). In muscle, glycogen provides fuel for muscle contraction. In contrast, liver
glycogen is largely converted to glucose that exits liver cells and enters the bloodstream
for transport to other tissues that require it. Both the mobilization and synthesis of
glycogen are regulated by hormones.

A. Regulation of Glycogen Phosphorylase

Glycogen phosphoryase is responsible for the breakdown of glycogen to produce
glucose 1 -phosphate. In muscle cells, glucose 1 -phosphate is converted to glucose
6 -phosphate that is used in glycolysis to produce ATP. In liver cells, glucose 6-phosphate
is hydrolyzed to free glucose that is secreted into the bloodstream where it can be taken
up by other tissues.

The activity of glycogen phosphorylase is regulated by several allosteric effectors
and by covalent modification (phosphorylation). Let’s take a few minutes to study the
regulation of glycogen phosphorylase because not only is it important in glycogen me-
tabolism, it’s also historically important.

The enzyme exists in four different forms as shown in Figure 12.22. The unphos-
phorylated form is called glycogen phosphorylase b (GPb) and the phosphorylated
form is called glycogen phosphorylase a (GPa). The enzyme is phosphorylated by a
kinase enzyme and dephosphorylated by a phosphatase.

Like other allosterically regulated enzymes, glycogen phosphorylase adopts two
conformations; the R conformation is the active conformation and the T conformation
is much less active. This is depicted in Figure 12.22 as a change in the shape of the cat-
alytic site: In the R conformation, inorganic phosphate (a substrate of the reaction) can
bind and in the T conformation binding of inorganic phosphate is inhibited.

Unphosphorylated GPb can exist in both inactive T conformations and active R
conformations. The allosteric site of the enzyme binds several effectors that cause a

12.6 Regulation of Glycogen Metabolism in Mammals 373

BOX 12.5 HEAD GROWTH AND TAIL GROWTH

Polymerization reactions can be described as either head
growth or tail growth. In a head growth mechanism, the
growing end of the chain is “activated” and cleavage of the
“high energy” linkage at the head of the molecule provides
the energy for the next addition of a monomer. In a tail growth
mechanism, the growing end does not contain the high energy
linkage; instead, the energy for the addition reaction comes
from the activated monomer.

Glycogen synthesis is an example of a tail growth
mechanism. The incoming monomer (UDP-glucose) is
activated and, when the reaction is complete, the end of the
glycogen chain is a simple hydroxyl group at the 4-carbon
atom of a glucose residue. DNA and RNA synthesis are also
examples of a tail growth mechanism. Protein synthesis
and fatty acid synthesis are examples of head growth
mechanism.

The differences between the two mechanisms become
clear when you think of the reverse reaction: degradation.

Head Growth

Glycogen and nucleic acids can be degraded by chopping off a
single residue. In the case of glycogen, synthesis and degrada-
tion are part of an ongoing process since the glycogen particle
serves as a storage molecule for glucose. In the case of nucleic
acids, especially DNA, the degradation reaction is an essential
part of DNA repair and proofreading that ensures DNA
replication is extremely accurate (Section 20.2C). Removal of
single residues does not prevent the polymer from serving im-
mediately as a substrate for further addition reactions.

Protein synthesis and fatty acid synthesis utilize head
growth mechanisms for synthesis. In this case, removal of an
end residue also removes the activated head so further addi-
tion reactions are not possible without an additional step to
“reactivate” the head. This is one reason why protein synthe-
sis errors cant be repaired and one reason why fatty acid
chains aren’t used as energy storage molecules in the same
way that glycogen is used.

Tail Growth

q Head

Tail

Head

Synthesis

Degradation

▲ Head and tail growth. In a head growth mechanism (left), incoming activated monomers are added to the “head” of the growing polymer.

(The end that contains the activated residue.) After the addition reaction, the polymer still contains an activated residue at the growing end. In
tail growth (right), the incoming activated monomer is added to the “tail” end of the growing polymer. The monomer substrate carries the energy
for its own addition reaction. When the polymer is degraded, a single residue is removed. Polymers that use a head growth mechanism will no
longer be a substrate for addition reactions following degradation because the activated head has been removed. Polymers that employ a tail
growth mechanism are still able to act as substrates for addition reactions.

shift in conformation. The allosteric site is close to the dimer interface between the two
monomers and both subunits change conformation simultaneously — a result that con-
forms to the concerted model of Monod, Wyman, and Changeux (Section 5.9C).

When ATP is bound, the activity of the enzyme is inhibited (T state). This is the nor-
mal state of activity since physiological concentrations of ATP are high and relatively con-
stant. When the AMP concentration rises, it displaces ATP from the allosteric site causing a
shift to the active R conformation and activation of glycogen breakdown. In muscle cells,
increasing AMP concentration results from strenuous muscle activity and signals the need
for more glucose 1 -phosphate to stimulate ATP production by glycolysis. The enzyme is in-
hibited by glucose 6-phosphate (feedback inhibition). There’s no need to continue glyco-
gen breakdown if glucose 6-phosphate concentration is sufficient to fuel glycolysis.

The main difference between the R conformation and the T conformation is the
position of a loop containing Asp-283 and nearby residues (the 280s loop). In the T
conformation, the negatively charged side chain of Asp-283 lies close to the pyridoxal
5-phosphate (PLP) cofactor at the catalytic site. This proximity prevents binding of
inorganic phosphate, inhibiting the reaction. In the R conformation, the position of this
loop shifts allowing inorganic phosphate to enter the active site.

374 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Glucose

▲ Figure 12.21

Degradation of glycogen. Glycogen phospho-
rylase catalyzes the phosphorolysis of glyco-
gen chains, stopping four residues from an
a-(l — * 6) branch point and producing one
molecule of glucose 1-phosphate for each
glucose residue mobilized. Further degrada-
tion is accomplished by the two activities of
the glycogen debranching enzyme. The
4-a-glucanotransferase activity catalyzes the
transfer of a trimer from a branch of the
limit dextrin to a free end of the glycogen
molecule. The amylo-l,6-glucosidase
activity catalyzes hydrolytic release of the
remaining a-{ 1 — » 6)-l i nked glucose residue.

Phosphofructokinase-1 (PFK-1) is
regulated in a similar manner by
ATP and AMP.

The structures of GPa and GPb are shown in Figure 12.23 in order to illustrate the
structural changes that take place when the enzyme is phosphorylated and dephospho-
rylated. The phosphoryl group is covalently attached to serine residue 14 (Ser-14) near
the N- terminal end of the protein.

In the unphosphorylated state (GPb), the N-terminal residues, including Ser-14,
associate with the surface near the catalytic site. In the phosphorylated state (GPa),
phosphoserine-14 interacts with two positively charged arginine residues near the
allosteric site. The remarkable shift in the location of the N-terminal end of the chain
cause other conformation changes in the enzyme; notably, a reorientation of two a
helices, the tower helices, on the other side of the dimer interface. This, in turn, affects
the position of the 280s loop controlling the transition between the active R conforma-
tion and the inactive T conformation.

The equilibrium between T and R is greatly shifted in favor of the R conformation
(active) when glycogen phosphorylase is phosphorylated (GPa). GPa is relatively
insensitive to ATP, AMP, and glucose 6-phosphate. In muscle cells, GPa will be formed
in response to hormones that signal the need for glucose and strenuous muscle activity.
This promotes rapid mobilization of glycogen. In liver cells, the liver version of glycogen
phosphorylase responds to the same hormones but in this case glycogen breakdown
leads to excretion of glucose that can be taken up by muscle cells. Liver glycogen phos-
phorylase a is inhibited by glucose by shifting GPa to the T conformation. This makes
sense since the presence of a high concentration of free glucose means that it’s not nec-
essary to continue producing glucose from glycogen.

The muscle version of glycogen phosphorylase is not inhibited by glucose since
muscle cells rarely see significant concentrations of free glucose. Muscle cells don’t con-
vert glucose 6-phosphate to glucose and any glucose taken up from the bloodstream is
quickly phosphoryated by hexokinase to glucose 6-phosphate.

Glycogen
phosphorylase b
(GPb)

Glycogen
phosphorylase a
(GPa)

Glucose

▲ Figure 12.22

Regulation of glycogen phosphorylase. Glycogen phosphorylase b is the unphosphorylated form of
the enzyme. Glycogen phosphorylase a is phosphorylated at a position near the allosteric site.
Phosphorylation is indicated by a purple ball at that site. The T conformation (red) is mostly
inactive and the R conformation (green) is active in glycogen breakdown as shown by binding of
inorganic phosphate (purple ball) to the catalytic site. The R conformation is greatly favored when
the enzyme is phosphorylated (glycogen phosphorylase a).

12.6 Regulation of Glycogen Metabolism in Mammals 375

T state

R state

Catalytic site
PLP'

Catalytic site
PLP

Ser-14' shift

Ser14-P'

Arg-14

Tower

helices

Catalytic site
PLP

▲ Figure 12.23

Phosphorylated and unphosphoylated forms of glycogen phosphorylase. PLP at the catalytic site is
shown as a space-filling molecule. The large shift in position of Ser-14 upon phospharylation to
Ser-14-P causes a conformational change that allows access to the catalytic site [PDB 3CEH, 1Z8D].

Gerty Cori and Carl Cori discovered in 1938 that glycogen phosphorylase activity
was regulated by AMP. Since then, glycogen phosphorylase has been one of the prime
examples of allosterically regulated enzymes, exciting three generations of biochemistry
students. Glycogen phosphorylase was the very first enzyme whose regulation by cova-
lent modification was demonstrated. Eddy Fischer and Edwin Krebs published their
result in 1956 and for a long time regulation by phosphorylation was thought to be an
unusual form of regulation confined to glycogen metabolism. Today, we know that
phosphorylation is a very common form of regulation in eukaryotes and it is the most
important part of many signal transduction pathways. There are hundreds of labs
studying signal transduction.

B. Hormones Regulate Glycogen Metabolism

Insulin, glucagon, and epinephrine are the principal hormones that control glycogen
metabolism in mammals. Insulin, a 51 -residue protein synthesized by the /3 cells of the
pancreas, is secreted when the concentration of glucose in the blood increases. High levels
of insulin are associated with the fed state of an animal. Insulin increases the rate of
glucose transport into muscle and adipose tissue via the GLUT4 glucose transporter
(Section 11.5A). Insulin also stimulates glycogen synthesis in the liver.

Glucagon, a peptide hormone containing 29 amino acid residues, is secreted by the
a cells of the pancreas in response to a low blood glucose concentration. Glucagon re-
stores the blood glucose concentration to a steady state level by stimulating glycogen

▲ Edmond (“Eddy”) H. Fischer (1920-)
(left) and Edwin G. Krebs (1918-2009)
(right) received the Nobel Prize in Physiology
or Medicine in 1992 “for their discoveries
concerning reversible protein phosphoryla-
tion as a biological regulatory mechanism.”

376

CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

degradation. Glucagon is extremely selective in its target because only liver cells are
rich in glucagon receptors. The effect of glucagon is opposite that of insulin and an ele-
vated glucagon concentration is associated with the fasted state.

The adrenal glands release the catecholamine epinephrine (also known as adrena-
line), in response to neural signals that trigger the fight or flight response (Figure 3.5c).
The epinephrine precursor, norepinephrine, also has hormone activity. Epinephrine
stimulates the breakdown of glycogen. It triggers a response to a sudden energy require-
ment whereas glucagon and insulin act over longer periods to maintain a relatively
constant concentration of glucose in the blood. Epinephrine binds to /3-adrenergic
receptors of liver and muscle cells and to a r adrenergic receptors of liver cells. The
binding of epinephrine to /3-adrenergic receptors or of glucagon to its receptors acti-
vates the adenylyl cyclase signaling pathway. The second messenger, cyclic AMP
(cAMP), then activates protein kinase A (PKA).

PKA phosphorylates a number of other proteins causing significant changes in me-
tabolism. Let’s look first at the regulation of glycogen metabolism by glucagon (Figure
12.24). When glucagon binds to its receptor it stimulates adenylate cyclase causing an in-
crease in cAMP that leads to activation of PKA. PKA phosphorylates glycogen synthase
converting the “a” form to the inactive cc b” form. This blocks glycogen synthesis. PKA also
phosphorylates another kinase called phosphorylase kinase. As the name implies, this is
the kinase that phosphorylates glycogen phosphorylase. PKA activates phosphorylase ki-
nase leading to conversion of glycogen phosphorylase b to the the active form, glycogen
phosphorylase a. The result is an increase in the rate of degradation of glycogen.

The net effect of glucagon (or epinephrine) is to block synthesis of glycogen and
stimulate its breakdown. The reciprocal regulation of these two enzymes is an impor-
tant feature of regulation in this pathway.

Glycogen synthase and glycogen phosphorylase are dephosphorylated by phospho-
protein phosphatase- 1, an enzyme that acts on many other substrates. As shown in
Figure 12.25, dephosphorylation leads to reciprocal inactivation of glycogen phospho-
rylase and activation of glycogen synthase. This results in synthesis of glycogen from
UDP- glucose and inhibition of glycogen breakdown. Insulin stimulates the activity of
phosphoprotein phosphatase- 1, thus causing the uptake of glucose into glycogen and its
depletion in the bloodstream. Prosphoprotein phosphatase- 1 also acts on phosphory-
lase kinase blocking further activation of glycogen phosphorylase.

C. Hormones Regulate Gluconeogenesis and Glycolysis

Now it’s time to return to our discussion of the regulation of gluconeogenesis and glycolysis.
Fructose 1,6-frzsphosphatase (FBPase) and phosphofructokinase- 1 (PFK-1) are the key
enzymes involved in the decision to either degrade glucose or synthesize it (Section 12.3).
Recall that these two enzymes are reciprocally regulated by the effector fructose
2,6-Hsphosphate (Figure 12.8). This effector molecule is synthesized from fructose
6-phosphate by phosphofructokinase-2 (PFK-2) and it is dephosphorylated back to fruc-
tose 6-phosphate by fructose 2,6-Z?zsphosphatase (F2,6BPase) (Figure 12.9). These two
enzymatic activities are located on the same bifunctional protein. The relationship among
the four enzymes and their products is summarized in Figure 12.26.

The F2,6BPase and PFK-2 activities in the bifunctional enzyme are regulated by
phosphorylation in a reciprocal manner. When the protein is phosphorylated, the
enyme acts as a fructose 2,6-Hsphosphatase and the phosphofructokinase activity is in-
hibited. Conversely, when the enzyme is unphosphorylated it acts as a phosphofructo-
kinase and the fructose 2,6-frisphosphatase activity is inhibited.

This is the same mode of reciprocal regulation we encountered with glycogen
phosphorylase and glycogen synthase, except this time the two enzyme activities are on
the same molecule. In the presence of glucagon, protein kinase A (PKA) is active and it
phosphorylates the bifunctional enzyme (Figure 12.27). Thus, glucagon stimulates glu-
coneogenesis and inhibits glycolysis in liver cells causing glucose levels in the blood-
stream to rise. At the same time, epinephrine can stimulate glycogen degradation and
inhibit glycogen synthesis in muscle cells. The result is more glucose for muscle cells
and more ATP from glycolysis.

12.6 Regulation of Glycogen Metabolism in Mammals 377

◄ Figure 12.24

Effects of glucagon on glycogen metabolism.

The binding of glucagon to its receptors
stimulates glycogen degradation via protein
kinase A.

ATP ADP

t Figure 12.25

Effect of insulin on glycogen metabolism. Insulin simulates the phosphatase activity of phosphoprotein
phosphatase-1, leading to inactivation of glycogen phosphorylase and activation of glycogen synthase.

ATP ATP ADP

Insulin

Phosphoprotein

phosphatase-1

378

CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Gluconeogenesis

Glycolysis

Figure 12.26 ▲

The role of fructose 2,6-Z;/sphosphate in
regulating glycolysis and gluconeogenesis.

12.7 Maintenance of Glucose Levels
in Mammals

Mammals maintain blood glucose levels within strict limits by regulating both the syn-
thesis and degradation of glucose. Glucose is the major metabolic fuel in the body.
Some tissues, such as brain, rely almost entirely on glucose for their energy needs. The
concentration of glucose in the blood seldom drops below 3 mM or exceeds 10 mM.
When the concentration of glucose in the blood falls below 2.5 mM, glucose uptake into
the brain is compromised, with severe consequences. Conversely, when blood glucose
levels are very high, glucose is filtered out of the blood by the kidneys accompanied by
osmotic loss of water and electrolytes.

The liver plays a unique role in energy metabolism participating in the intercon-
versions of all types of metabolic fuels: carbohydrates, amino acids, and fatty acids.

Figure 12.27 ►

Effect of glucagon on gluconeogenesis.

Glucagon binds to its receptor, causing acti-
vation of adenylate cyclase. Increased levels
of cAMP activate protein kinase A, which
phosphorylates the bifunctional enzyme
leading to activation of fructose 2,6-
b/sphosphatase activity. In the absence of
the effector fructose 2,6-b/sphosphate, fruc-
tose l,6-b/'sphosphatase is activated and
this increases flux in the gluconeogenesis
pathway.

F2,6P

12.7 Maintenance of Glucose Levels in Mammals 379

Anatomically, the liver is centrally located in the circulatory system (Figure 12.28).
Most tissues are perfused in parallel with the arterial system supplying oxygenated
blood and the venous circulation returning blood to the lungs for oxygenation. The
liver, however, is perfused in series with the visceral tissues (gastrointestinal tract, pan-
creas, spleen, and adipose tissue); blood from these tissues drains into the portal vein
and then flows to the liver. This means that after the products of digestion are absorbed
by the intestine, they pass immediately to the liver. Using its specialized complement of
enzymes, the liver regulates the distribution of dietary fuels and supplies fuel from its
own reserves when dietary supplies are exhausted.

The consumption of glucose by tissues removes dietary glucose from the blood.
When glucose levels fall, liver glycogen and gluconeogenesis become the sources of glu-
cose. However, since these sources are limited, hormones act to restrict the use of glucose
to those cells and tissues that absolutely depend on glycolysis for generating ATP (kidney
medulla, retina, red blood cells, and parts of the brain). Other tissues can generate ATP
by oxidizing fatty acids mobilized from adipose tissue (Sections 16. 1C and 16.2).

The complexity of carbohydrate metabolism in mammals is evident from the
changes that occur on feeding and starvation. In the 1960s, George Cahill examined the
glucose utilization of obese patients as they underwent therapeutic starvation. After an
initial feeding of glucose, the subjects received only water, vitamins, and minerals.
Cahill noted that glucose homeostasis (maintenance of constant levels in the circula-
tion) proceeds through five phases. Figure 12.29, based on Cahill’s observations, sum-
marizes the metabolic changes in the five phases.

1. During the initial absorptive phase (the first four hours), dietary glucose enters the
liver via the portal vein and most tissues use glucose as the primary fuel. Under
these conditions, the pancreas secretes insulin, which stimulates glucose uptake by
muscle and adipose tissue via GLUT4. The glucose taken up by these tissues is

◄ Figure 12.28

Placement of the liver in the circulatory system.

Most tissues are perfused in parallel. How-
ever, the liver is perfused in series with vis-
ceral tissues. Blood that drains from the in-
testine and other visceral tissues flows to the
liver via the portal vein. The liver is therefore
ideally placed to regulate the passage of
fuels to other tissues.

380 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Figure 12.29 ►

Five phases of glucose homeostasis. The graph,
based on observations of a number of individ-
uals, illustrates glucose utilization in a 70 kg
man who consumed 100 g of glucose and
then fasted for 40 days.

1 2 3 4 5

The effect of insulin and diabetes on
the production of ketone bodies is
described in Section 16.11 (Box 16.6).

phosphorylated to glucose 6-phosphate, which cannot diffuse out of the cells. Liver
cells also absorb glucose and convert it to glucose 6-phosphate. Excess glucose is
stored as glycogen in liver and muscle cells.

2. When the dietary glucose is consumed, the body mobilizes liver glycogen to maintain
circulating glucose levels. In the liver, glucose 6-phosphatase catalyzes the hydroly-
sis of glucose 6-phosphate to glucose, which exits the liver via glucose transporters.
Glycogen in muscle (which lacks glucose 6-phosphatase) is metabolized to lactate
to produce ATP for contraction; the lactate is used by other tissues as a fuel or by
the liver for gluconeogenesis.

3. After about 24 hours, liver glycogen is depleted, and the only source of circulating
glucose is gluconeogenesis in the liver, using lactate, glycerol, and alanine as precur-
sors. Fatty acids mobilized from adipose tissue become an alternate fuel for most
tissues. The obligatory glycolytic tissues continue to use glucose and produce lac-
tate, which is converted to glucose in the liver by the Cori cycle; this cycle makes
energy, not carbon, from fatty acid oxidation in the liver available to other tissues.

4. Gluconeogenesis in the liver continues at a high rate for a few days, then decreases.
As starvation progresses, gluconeogenesis in the kidney becomes proportionately
more significant. Proteins in peripheral tissues are broken down to provide gluco-
neogenic precursors. In this phase, the body adapts to several alternate fuels.

5. In prolonged starvation, there is less gluconeogenesis and lipid stores are depleted.
If refeeding does not occur, death will follow. On refeeding, metabolism is quickly
restored to the conditions of the fed state.

We have seen how glucose, a major fuel, can be stored in polysaccharide form and
mobilized as needed. Glucose can also be synthesized from noncarbohydrate precursors
by the reactions of gluconeogenesis. We have seen that glucose can be oxidized by the pen-
tose phosphate pathway to produce NADPH or transformed by glycolysis into pyruvate.

Diabetes mellitus (DM) is a metabolic disease that results from improper regula-
tion of carbohydrate and lipid metabolism. Despite an ample supply of glucose, the
body behaves as though starved and glucose is overproduced by the liver and underused
by other tissues. As a result, the concentration of glucose in the blood is extremely high.
The levels of glucose in the blood often exceed the capacity of the kidney to reabsorb
glucose so some of it spills into the urine. The high concentration of glucose in urine
draws water osmotically from the body.

There are two types of diabetes both of which arise from faulty control of fuel
metabolism by the hormone insulin. In Type 1 diabetes mellitus (also called insulin-
dependent diabetes mellitus, or IDDM) damage to the /3 cells of the pancreas, where
insulin is synthesized, results in diminished or absent secretion of insulin. This au-
toimmune disease is characterized by early onset (usually before age 15). Patients are
thin and exhibit hyperglycemia (high blood glucose levels), dehydration, excessive
urination, hunger, and thirst. In Type 2 (also called non-insulin-dependent diabetes,
or NIDDM), chronic hyperglycemia results from insulin resistance — decreased

12.8 Glycogen Storage Diseases 381

sensitivity to insulin possibly caused by a shortage or decreased activity of insulin re-
ceptors. Insulin secretion may be normal and circulating levels of insulin may even
be elevated. This type is also known as adult- onset diabetes (although its incidence is
increasing among children) and it is usually associated with obesity. Type 2 diabetes
affects about 5% of the population and Type 1 affects about 1%. In addition, about
2% to 5% of pregnant women develop a form of diabetes. Most women who exhibit
gestational diabetes return to normal after giving birth but are at risk for developing
Type 2 diabetes.

To understand diabetes, we must consider the functions of insulin. Insulin stimu-
lates the synthesis of glycogen, triacylglycerols, and proteins and inhibits the breakdown
of these compounds. Insulin also stimulates glucose transport into muscle cells and
adipocytes. When insulin levels are low in IDDM, glycogen is broken down in the liver
and gluconeogenesis occurs regardless of the glucose supply. In addition, glucose uptake
and its use in peripheral tissues are restricted.

12.8 Glycogen Storage Diseases

Several metabolic diseases are related to the storage of glycogen. The general rule about
metabolic diseases is that they usually affect the activity of nonessential genes and en-
zymes. Defects in essential genes are usually lethal and don’t show up as metabolic
diseases.

Many metabolic enzymes in humans are encoded by gene families. Different ver-
sions are expressed in different tissues. In the case of enzymes involved in glycogen me-
tabolism, the most common versions are found in liver and muscle. A deficiency in one
of these enzymes will produce severe symptoms but may not be lethal. There are nine
types of glycogen storage diseases resulting from defects in glycogen metabolism.

Type 0: In type 0a, the activity of liver glycogen synthase is affected. The gene for this
enzyme is on the short arm of chromosome 12 at locus 12pl2.2 (MIM = 240600).
This is a severe disease causing early death in cases where the activity is very low. Type
Ob affects the muscle version of glycogen synthase whose gene is on the long arm of
chromosome 19 at 19ql3.3 (MIM = 611556). Patients have no muscle glycogen
and are unable to engage in strenuous physical activity.

Type I: The most common glycogen storage disease is called von Gierke disease. It
is caused by a deficiency in glucose 6-phosphatase (Type la, MIM = 23220) whose
gene is on chromosome 17 (17q21). Defects in the complex that transports glucose
across the endoplasmic reticulum (Section 21. ID) also cause von Gierke’s disease.
Type lb affects the glucose 6-phosphate transporter (chromosome 11 (llq23),
MIM = 232220) and type lc affects the phosphate transporter (chromosome 6
(6p21.3), MIM = 232240). Patients are unable to secrete glucose leading to accu-
mulation of glycogen in the liver and kidneys.

Type II: Patients suffering from type II disease, known as Pompe’s disease, suffer
from reduced activity of a-l,4-glucosidase, or acid maltase, an enzyme required for
glycogen breakdown in lysozomes (MIM = 232300). The gene is on chromosome
17 (17q25.2). The defect causes glycogen to accumulate in lysosomes leading to
problems with muscle tissue, especially in the heart. In the most severe forms,
children die within the first few years of life.

Type III: Type III is Cori disease, characterized by defects in the gene encoding the
glycogen debranching enzyme in liver and muscle (chromosome 1 (lp21), MIM =
232400). People suffering from this disease have weakened muscles because they
are unable to mobilize all of the stored glycogen. Some defects have very mild
symptoms.

Type IV: Often called Anderson’s disease, the mutations occur in the gene for liver
branching enzyme found on chromosome 3 (3pl2, MIM = 232500). Long-chain
polysaccharides accumulate in patients with these mutations, resulting in death
within a few years from heart failure or liver failure.

MIM numbers refer to the Online
Mendelian Inheritance in Man (0MIM)
database at: ncbi.nlm.nih.gov/omim

382

CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Type V: McAr die’s disease (type V glycogen storage disease) is caused by a
deficiency of muscle glycogen phosphorylase (MIM = 232600). The gene is on
chromosome 11 (llql3). Individuals having this genetic disease cannot perform
strenuous exercise and suffer painful muscle cramps.

Type VI: Hers’ disease (type VI) is a mild form of glycogen storage disease due to a
deficiency in liver glycogen phosphorylase (MIM = 232700). Several mutant alleles
interfere with proper splicing of the primary transcript from the gene on chromo-
some 14 (14q21).

Type VII: Mutations in the gene for muscle phosphofructokinase- 1 cause Tarui’s
disease, characterized by inability to exercise and muscle cramps (MIM = 232800).
The gene for this isozyme is on chromosome 12 (12ql3.3).

Type VIII: Now recognized as a subtype of type IX.

Type IX: This form of glycogen storage disease manifests as muscle weakness
and/or muscle cramps. The symptoms are usually mild. All subtypes are due to
mutations in the genes for the various subunits of glycogen phosphorylase kinase.
Type IXa: liver a subunit gene on the X chromosome at Xp20 (MIM = 300798).
Type IXb: /3 subunit gene at 16ql2 (MIM = 172490). Type IXc: liver y subunit gene
at 16pl2 (MIM = 172471). Type IXd: muscle a subunit gene on the X chromosome
at Xql3 (MIM = 311870).

Summary

1. Gluconeogenesis is the pathway for glucose synthesis from non-
carbohydrate precursors. The seven near- equilibrium reactions of
glycolysis proceed in the reverse direction in gluconeogenesis.
Four enzymes specific to gluconeogenesis catalyze reactions that
bypass the three metabolically irreversible reactions of glycolysis.

2. Noncarbohydrate precursors of glucose include pyruvate, lactate,
alanine, and glycerol.

3. Gluconeogenesis is regulated by glucagon, allosteric modulators,
and the concentrations of its substrates.

4. The pentose phosphate pathway metabolizes glucose 6-phosphate
to generate NADPH and ribose 5-phosphate. The oxidative stage
of the pathway generates two molecules of NADPH per molecule
of glucose 6-phosphate converted to ribulose 5 -phosphate and
C0 2 . The nonoxidative stage includes isomerization of ribulose
5-phosphate to ribose 5-phosphate. Further metabolism of pen-
tose phosphate molecules can convert them to glycolytic interme-
diates.The combined activities of transketolase and transaldolase

convert pentose phosphates to triose phosphates and hexose
phosphates.

5. Glycogen synthesis is catalyzed by glycogen synthase, using a
glycogen primer and UDP- glucose.

6. Glucose residues are mobilized from glycogen by the action of
glycogen phosphorylase. Glucose 1 -phosphate is then converted
to glucose 6-phosphate.

7. Glycogen degradation and glycogen synthesis are reciprocally reg-
ulated by hormones. Kinases and phosphatases control the activi-
ties of the interconvertible enzymes glycogen phosphorylase and
glycogen synthase.

8. Mammals maintain a nearly constant concentration of glucose in
the blood. The liver regulates the amount of glucose supplied by
the diet, glycogenolysis, and other fuels.

9. Glycogen storage diseases result from defects in genes required for
glycogen metabolism.

Problems

1. Write a balanced equation for the synthesis of glucose from pyru-
vate. Assuming that the oxidation of NADH is equal to 2.5 ATP
equivalents (Section 14.11), how many ATP equivalents are re-
quired in this pathway? Convert this to kj mol 1 and explain how
this value compares to the total energy required to synthesize
glucose from C0 2 and H 2 0.

2. What important products of the citric acid cycle are required for
gluconeogenesis from pyruvate?

3. Epinephrine promotes the utilization of stored glycogen for gly-
colysis and ATP production in muscles. How does epinephrine
promote the use of liver glycogen stores for generating the energy
needed by contracting muscles?

4. (a) In muscle cells, insulin stimulates a protein kinase that cat-

alyzes phosphorylation of protein phosphatase- 1, thereby
activating it. How does this affect glycogen synthesis and
degradation in muscle cells?

(b) Why does glucagon selectively regulate enzymes in the liver
but not in other tissues?

5. The polypeptide hormone glucagon is released from the pancreas
in response to low blood glucose levels. In liver cells, glucagon
plays a major role in regulating the rates of the opposing glycolysis

Selected Readings 383

and gluconeogenesis pathways by influencing the concentrations
of fructose 2,6-frzsphosphate (F2,6 BP). If glucagon causes a de-
crease in the concentrations of F2,6 BP, how does this result in an
increase in blood glucose levels?

6. When the concentration of glucagon rises in the blood, which of
the following enzyme activities is decreased? Explain.

Adenylyl cyclase
Protein kinase A
PFK-2 (kinase activity)

Fructose 1, 6-Hsphosphatase

7. (a) Is the energy required to synthesize glycogen from glucose

6-phosphate greater than the energy obtained when glycogen
is degraded to glucose 6-phosphate?

(b) During exercise, glycogen in both muscle and liver cells can
be converted to glucose metabolites for ATP generation in
the muscles. Do liver glycogen and muscle glycogen supply
the same amount of ATP to the muscles?

8. Individuals with a total deficiency of muscle glycogen phosphory-
lase (McArdle s disease) cannot exercise strenuously due to mus-
cular cramping. Exertion in these patients leads to a much greater
than normal increase in cellular ADP and Pj. Furthermore, lactic
acid does not accumulate in the muscles of these patients, as it
does in normal individuals. Explain the chemical imbalances in
McArdle’s disease.

9. Compare the number of ATP equivalents generated in the break-
down of one molecule of glucose 1 -phosphate into two molecules
of lactate with the number of ATP equivalents required for the
synthesis of one molecule of glucose 1 -phosphate from two mole-
cules of lactate. (Assume anaerobic conditions.)

10 . (a) How does the glucose-alanine cycle allow muscle pyruvate to
be used for liver gluconeogenesis and subsequently returned
to muscles as glucose?

(b) Does the glucose-alanine cycle ultimately provide more en-
ergy for muscles than the Cori cycle does?

11. Among other effects, insulin is a positive modulator of the en-
zyme glucokinase in liver cells. If patients with diabetes mellitus
produce insufficient insulin, explain why these patients cannot
properly respond to increases in blood glucose.

12. Glycogen storage diseases (GSDs) due to specific enzyme defi-
ciencies can affect the balance between glycogen stores and blood
glucose. Given the following diseases, predict the effects of each
on (1) the amount of liver glycogen stored and (2) blood glucose
levels.

(a) Von Gierke disease (GSD-la), defective enzyme: glucose
6-phosphatase.

(b) Cori’s disease (GSD III), defective enzyme: amylo-1,6 glu-
cosidase (debranching enzyme).

13 . The pentose phosphate pathway and the glycolytic pathway are
interdependent, since they have in common several metabolites
whose concentrations affect the rates of enzymes in both path-
ways. Which metabolites are common to both pathways?

14 . In many tissues, one of the earliest responses to cellular injury is a
rapid increase in the levels of enzymes in the pentose phosphate
pathway. Ten days after an injury, heart tissue has levels of glucose
6-phosphate dehydrogenase and 6-phosphogluconate dehydroge-
nase that are 20 to 30 times higher than normal, whereas the levels
of glycolytic enzymes are only 10% to 20% of normal. Suggest an
explanation for this phenomenon.

15 . (a) Draw the structures of the reactants and products for the

second reaction catalyzed by transketolase in the pentose
phosphate pathway. Show which carbons are transferred.

(b) When 2- [ 14 C] -glucose 6-phosphate enters the pathway,
which atom of fructose 6-phosphate produced by the reac-
tion in Part (a) is labeled?

Selected Readings

Gluconeogenesis

Hanson, R. W., and Hakimi, P. (2008). Born to
run. Biochimie. 90:838-842.

Hanson, R. W., and Reshef, L. (1997). Regulation
of phosphenolpyruvate carboxykinase (GTP) gene
expression. Annu. Rev. Biochem. 66:581-611.
Describes the metabolic control of gene expression.

Hines, J. K., Chen, X., Nix, J. C., Fromm, H. J., and
Honzatko, R. B. (2007). Structures of mammalian
and bacterial fructose- 1,6-bisphosphatase reveal the
basis for synergism in AMP/fructose 2,6-bisphos-
phate inhibition./. Biol Chem. 282:36121-36131.

Jitrapakdee, S., and Wallace, J. C. (1999). Struc-
ture, function and regulation of pyruvate carboxy-
lase. Biochem. J. 340:1-16.

Kemp, R. G. and Gunasekera, D. (2002). Evolution
of the allosteric ligand sites of mammalian
phosphofructo- 1 -kinase. Biochem. Biochemistry
41:9426-9430.

Ou, X., Ji, C., Han, X., Zhao, X., Li, X., Mao, Y.,
Wong, L-L., Bartlam, M., and Rao, Z. (2006).

Crystal structure of human glycerol 3 -phosphate
dehydrogenase (GPD1 )./. Mol. Biol. 357:858-869.

Pilkis, S. J., and Granner, D. K. (1992). Molecular
physiology of the regulation of hepatic gluconeo-
genesis and glycolysis. Annu. Rev. Physiol.
57:885-909.

Rothman, D. L., Magnusson, I., Katz, L. D.,
Shulman, R. G., and Shulman, G. I. (1991).
Quantitation of hepatic glycogenolysis and gluco-
neogenesis in fasting humans with 13 C NMR.
Science. 254:573-576. Describes the continuous
operation of the pathway of gluconeogenesis in
humans.

Sullivan, S. M., and Holyoak (2008). Enzymes with
lid-gated active sites must operate by an induced fit
mechanism instead of conformational selection.
Proc. Natl. Acad. Sci. (USA) 105:13829-13834.

van de Werve, G., Lange, A., Newgard, C., Mechin,
M.-C., Li, Y., and Berteloot, A. (2000). New lessons
in the regulation of glucose metabolism taught by
the glucose 6-phosphatase system. Eur. J. Biochem.
267:1533-1549. Explains why there is still much to

learn about the catalytic site and the transporter
associated with this enzyme.

Xue, Y., Huang, S., Liang, J. Y., Zhang, Y., and
Lipscomb, W. N. (1994). Crystal structure of
fructose- 1,6-bisphosphatase complexed with
fructose 2,6-bisphosphate, AMP, and Zn2+ at 2.0- A
resolution: aspects of synergism between inhibitors.
Proc. Natl. Acad. Sci. (USA) 91:12482-12486.

Pentose Phosphate Pathway

Au, S.W.N., Gover, S., Lam, V.M.S., and Adams,
M.J. (2000) Human glucose-6-phospate dehydro-
genase: the crystal structure reveals a structural
NADP + molecule and provides insights into en-
zyme deficiency. Structure 8:293-303.

Wood, T. (1985). The Pentose Phosphate Pathway.
(Orlando: Academic Press).

Wood, T. (1986). Physiological functions of the
pentose phosphate pathway. Cell Biochem. Pune.
4:241-247.

384 CHAPTER 12 Gluconeogenesis, The Pentose Phosphate Pathway, and Glycogen Metabolism

Glycogen Metabolism

Barford, D. Hu, S-H., and Johnson, L. N. (1991).
Structural mechanisms for glycogen phosphorylase
control by phosphorylation and AMR /. Mol. Biol
218:233-260.

Chou, J. Y., Matern, D., Mansfield, B. C., and Chen,
Y. T. (2002). Type I glycogen storage diseases: dis-
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Mol Med. 2:121-143.

Cohen, P., Alessi, D. R., and Cross, D. A. E. (1997).
PDK1, one of the missing links in insulin signal
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Fischer, E. (2010). Memories of Ed Krebs. /. Biol.
Chem. 285:4267.

Johnson, L. N. (2009). Novartis Medal Lecture:

The regulation of protein phosphorylation.
Biochem. Soc. Trans. 37:627-641.

Johnson, L. N., and Barford, D. (1990). Glycogen
phosphorylase: the structural basis of the allosteric

response and comparison with other allosteric
proteins./. Biol. Chem. 265:2409-2412.

Johnson, L. N., Lowe, E. D., Noble, M. E. M., and
Owen, D. J. (1998). The structural basis for sub-
strate recognition and control by protein kinases.
FEBS Lett. 430:1-11.

Larner, J. (1990). Insulin and the stimulation of
glycogen synthesis: the road from glycogen synthase
to cyclic AMP- dependent protein kinase to insulin
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Melendez-Hevia, E., Waddell, T. G., and Shelton,

E. D. (1993). Optimization of molecular design in
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Murray, R. K., Bender, D. A., Kennelly, P. J., Rodwell,
V. W., and Weil. P. A. (2009). Harpers Illustrated
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Pinotsis, N., Leonidas, D. D., Chrysina, E. D.,
Oikonomakos, N. G., and Mavridis, I. M.

(2003). The binding of f3- and y-cyclodextrins
to glycogen phosphorylase b: kinetic and
crystallographic studies. Prot. Sci.

12:1914-1924.

Shepherd, P. R., Withers, D. J., and Siddle, K.
(1998). Phosphoinositide 3-kinase: the key switch
mechanism in insulin signalling. Biochem. J.
333:471-490.

Smythe, C., and Cohen, P. (1991). The discovery
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glycogen biosynthesis. Eur. J. Biochem.
200:625-631.

Villar-Palasi, C., and Guinovart, J. J. (1997). The
role of glucose 6-phosphate in the control of
glycogen synthase. FASEB J. 11:544-558.

o c

_ o

° o o o

° o

o o

° c

o o

The Citric Acid Cycle

I n the last two chapters we were mainly concerned with the synthesis and degradation
of complex carbohydrates such as glucose. We saw that the biosynthetic pathway
leading to glucose began with pyruvate and oxaloacetate and that pyruvate was the
end product of glycolysis. In this chapter we will describe pathways that interconvert a
number of simple organic acids. Several of these compounds are essential precursors
for the biosynthesis of amino acids, fatty acids, and porphyrins.

Acetyl CoA is one of the key intermediates in the interconversion of small organic
acids. Acetyl CoA is formed by the oxidative decarboxylation of pyruvate with the re-
lease of C0 2 . This reaction is catalyzed by pyruvate dehydrogenase, an enzyme that we
briefly encountered in Section 11.3 when we discussed the fate of pyruvate. We begin
this chapter with a more detailed description of this important enzyme.

The acetyl group (a two-carbon organic acid) from acetyl CoA can be transferred to
the four-carbon dicarboxylic acid, oxaloacetate, to form a new six-carbon tricarboxylic
acid known as citrate (citric acid). Citrate can then be oxidized in a seven-step pathway
to regenerate oxaloacetate and release two molecules of C0 2 . Oxaloacetate can recom-
bine with another molecule of acetyl CoA and the citrate oxidation reactions are re-
peated. The net effect of this eight- enzyme cyclic pathway is the complete oxidation of an
acetyl group to C0 2 and the transfer of electrons to several cofactors to form reducing
equivalents. The pathway is known as the citric acid cycle, the tricarboxylic acid cycle
(TCA cycle), or the Krebs cycle, after Hans Krebs who discovered it in the 1930s.

The citric acid cycle lies at the hub of energy metabolism in eukaryotic cells —
especially in animals. The energy released in the oxidations of the citric acid cycle is
largely conserved as reducing power when the coenzymes NAD® and ubiquinone (Q)
are reduced to form NADH and QH 2 . This energy is ultimately derived from pyruvate
(via acetyl CoA). Since pyruvate is the end product of glycolysis, we can think of the
citric acid cycle as a series of reactions that complete the oxidation of glucose. NADH
and QH 2 are substrates in the reactions of membrane-associated electron transport
leading to the formation of a proton gradient that drives the synthesis of ATP (Chapter 14).

Since citric acid reacts catalytically
in the tissue it is probable that it is
removed by a primary reaction but
regenerated by a subsequent reac-
tion. In the balance sheet no citrate
disappears and no intermediate
products accumulate.

— H. A. Krebs and
W. A. Johnson (1937)

Top: Citrate synthase with its product citrate in the active site. This enzyme catalyzes the first step of the citric acid cycle.
[PDB 1CTS]

385

386

CHAPTER 13 The Citric Acid Cycle

Hans Krebs and W. A. Johnson proposed the citric acid cycle in 1937 in order to ex-
plain several puzzling observations. They were interested in understanding how the oxida-
tion of glucose in muscle cells was coupled to the uptake of oxygen. Albert Szent-Gyorgyi
had previously discovered that adding a four-carbon dicarboxylic acid — succinate, fumarate,
or oxaloacetate — to a suspension of minced muscle stimulated the consumption of 0 2 .
The substrate of the oxidation was carbohydrate, either glucose or glycogen. Especially in-
triguing was the observation that adding small amounts of four-carbon dicarboxylic acids
caused larger amounts of oxygen to be consumed than were required for their own oxi-
dation. This indicated that these four-carbon organic acids had catalytic effects.

Krebs and Johnson observed that citrate, a six-carbon tricarboxylic acid, and
a-ketoglutarate, a five-carbon compound, also had a catalytic effect on the uptake of
0 2 . They proposed that citrate was formed from a four-carbon intermediate and an un-
known two-carbon derivative of glucose (later shown to be acetyl CoA). The cyclic
nature of the pathway explained how its intermediates could act catalytically without
being consumed. Albert Szent-Gyorgyi received the Nobel Prize in Physiology or Medicine
in 1937 for his work on respiration, including the catalytic role of fumarate in biological
combustion processes. Hans Krebs was awarded the Nobel Prize in Physiology or Medicine
in 1953 for discovering the citric acid cycle.

In muscle cells, the intermediates in the citric acid cycle are almost exclusively used
in the cyclic pathway of energy metabolism. In these cells, the metabolic machinery is
mainly devoted to extracting energy from glucose in the form of ATP. This is why it was
possible to recognize the cyclic nature of the pathway by carrying out experiments on
muscle extracts. In other cells, the intermediates of the citric acid cycle are the starting
points for many biosynthetic pathways. Thus, the enzymes of the citric acid cycle play a
key role in both anabolic and catabolic reactions.

Many of these same enzymes are found in prokaryotes although few bacteria pos-
sess a complete citric acid cycle. In this chapter, we examine the reactions of the citric
acid cycle as they occur in eukaryotic cells. We will explore how these enzymes are regu-
lated. Next we will introduce the various biosynthetic pathways that require citric acid
cycle intermediates and examine the relationship of these pathways to the main reac-
tions of the cyclic pathway in eukaryotes and the partial pathways in bacteria. We will
also look at pathways involving glyoxylate, specifically the glyoxylate shunt and the gly-
oxylate cycle. These are pathways that are closely related to the citric acid cycle. Finally,
we will discuss the evolution of the citric acid cycle enzymes.

BOX 13.1 AN EGREGIOUS ERROR

In 1937, Krebs and Johnson submitted a
paper to Nature outlining their discov-
ery of citric acid as a catalyst in the oxi-
dation of glucose by muscle tissue. The
journal declined to publish the paper on
the grounds that it had too many papers
in press. Krebs writes in his memoirs,
“This was the first time in my career,
after having published more than fifty
papers, that I experienced a rejection or
semi-rejection.”

Krebs and Johnson published the
paper in the journal Enzymologia and

Krebs went on to win the Nobel Prize
based largely on this paper. It took
Nature 51 years to publically recognize
the mistake it made. An editor wrote in
the October 28, 1988 issue, “An editor’s
nightmare is to reject a Nobel-prizewin-
ning paper. . . . Rejection of Hans Krebs’
discovery of the tricarboxylic (or
Krebs’) cycle, a pivot of biochemical
metabolism, remains Nature's most
egregious error (as far as we know).”

► Hans Krebs (1900-1981). Krebs was awarded the
Nobel Prize in Physiology or Medicine in 1953 for
his discovery of the citric acid cycle. He is shown
here beside a Warburg apparatus for measuring
oxygen consumption in metabolizing tissue. Krebs
worked with Otto Warburg in the 1920s.

13.1 Conversion of Pyruvate to Acetyl CoA 387

13.1 Conversion of Pyruvate to Acetyl CoA

Pyruvate is a key substrate in a number of reactions, as described in Section 1 1.3. In this
chapter we are concerned with the conversion of pyruvate to acetyl CoA since acetyl
CoA is the main substrate of the citric acid cycle. The reaction is catalyzed by a large
complex of enzymes and cofactors known as the pyruvate dehydrogenase complex
(Figure 13.1). The stoichiometry of the complete reaction is

coo°

S-CoA

Pyruvate

< r — ° + HS-CoA + NAD©

dehydrogenase
>

^ ° + C0 2 + NADH

ch 3

Pyruvate

Acetyl CoA

▲ Figure 13.1

Electron micrograph of pyruvate dehydrogenase
complexes from E. coli.

where HS-CoA is coenzyme A. This is the first step in the oxidation of pyruvate and the
products of the reaction are acetyl CoA, one molecule of carbon dioxide, and one mole-
cule of reducing equivalent (NADH). The pyruvate dehydrogenase reaction is an oxida-
tion-reduction reaction. In this case, the oxidation of pyruvate to C0 2 is coupled to the
reduction of NAD® to NADH. The net result is the transfer of two electrons from
pyruvate to NADH.

The pyruvate dehydrogenase complex is a multienzyme complex containing multiple
copies of three distinct enzymatic activities: pyruvate dehydrogenase (E x subunits),
dihydrolipoamide acetyltransferase (E 2 subunits), and dihydrolipoamide dehydroge-
nase (E 3 subunits). The oxidative decarboxylation of pyruvate can be broken down into
five steps. (In each step of the following reactions the fates of the atoms from pyruvate
are shown in red.)

1. The E : component contains the prosthetic group thiamine diphosphate (TDP). As
we saw in Chapter 7, TDP (vitamin B x ) plays a catalytic role in a number of decar-
boxylase reactions. The initial reaction results in the formation of a hydroxyethyl-
TDP intermediate and the release of C0 2 .

The systematic names of the enzymes
in the complex are: pyruvate lipoamide
2-oxidoreductase (E^; acetyl CoA:dihy-
drolipoamide S-acetyltransferase
(E 2 ); and dihydrolipoamide:NAD©
oxidoreductase (E 3 ).

H,C

R— N x S + H 3 C — C — COO° + H @

h 3 c

Thiamine

diphosphate

(TDP)

Pyruvate

— 7 > R— N x S + C0 2

Pyruvate

dehydrogenase

HoC — C— OH

Hydroxyethylthiamine
diphosphate (HETDP)

(13.2)

Note that the reactive form of TDP is the carbanion or ylid form. The carbanion form
is relatively stable because of the unique environment of the coenzyme bound to the
protein (Section 7.6). The product of the first step is the carbanion form of hydrox-
yethyl-TDP. The mechanism is similar to the pyruvate decarboxylase mechanism
(Section 7.7).

2. In the second step, the two -carbon hydroxylethyl group is transferred to the
lipoamide group of E 2 . The lipoamide group consists of lipoic acid covalently
bound by an amide linkage to a lysine residue of an E 2 subunit (Figure 7.29). This
particular coenzyme is only found in pyruvate dehydrogenase and related enzymes.

388

CHAPTER 13 The Citric Acid Cycle

The transfer reaction is catalyzed by the E x component of the pyruvate dehydroge-
nase complex.

Lipoamide

H 3 C — C — OH

HETDP

Ylid Acetyl-dihydrolipoamide

(13.3)

In this reaction, the oxidation of hydroxyethyl-TDP is coupled to the reduction of
the disulfide of lipoamide and the acetyl group is transferred to one of the
sulfhydryl groups of the coenzyme regenerating the ylid form of TDP.

3. The third step involves the transfer of the acetyl group to HS-CoA, forming acetyl
CoA and leaving the lipoamide in the reduced dithiol form. This reaction is cat-
alyzed by the E 2 component of the complex.

H 3 C-

e 2

HS-CoA

H 3 C — c — S-CoA

Acetyl-dihydrolipoamide

Dihydrolipoamide Acetyl CoA

(13.4)

4. The reduced lipoamide of E 2 must be reoxidized in order to regenerate the pros-
thetic group for additional reactions. This is accomplished in step 4 by transferring
two protons and two electrons from the dithiol form of lipoamide to FAD. FAD is
the prosthetic group of E 3 and the redox reaction produces the reduced coenzyme
(FADH 2 ). (Recall from Section 7.5 that FADH 2 carries two electrons and two pro-
tons that are usually acquired as a single proton and a hydride ion.)

Dihydrolipoamide Lipoamide

(13.5)

5. In the final step, E 3 -FADH 2 is reoxidized to FAD. This reaction is coupled to the
reduction of NAD © .

E 3 — FADH 2 + NAD© > E 3 — FAD + NADH + H© (13.6)

The oxidation of E 3 -FADH 2 regenerates the original pyruvate dehydrogenase com-
plex, completing the catalytic cycle. Step 5 produces NADH and H©. Note that one
proton is released in step 5 and one proton is taken up in step 1 so that the overall
stoichiometry of the pyruvate dehydrogenase reaction shows no net gain or loss of
protons (Reaction 13.1).

The interplay of five coenzymes in the pyruvate dehydrogenase complex illustrates the
importance of coenzymes in metabolic reactions. Two of the coenzymes are cosubstrates
(HS-CoA and NAD©), and three are prosthetic groups (TDP, lipoamide, and FAD — one

13.1 Conversion of Pyruvate to Acetyl CoA

389

cofactor is bound to each type of subunit). The lipoamide groups bound to E 2 are prima-
rily responsible for transferring reactants from one active site in the complex to another. A
lipoamide picks up a two-carbon unit from hydroxyethyl-TDP in step 2 to form the
acetyl-dihydrolipoamide intermediate. This intermediate is repositioned in the active site
of dihydrolipoamide acetyltransferase where the two- carbon group is transferred to coen-
zyme A in step 3. The reduced lipoamide produced in that reaction is then moved to the ac-
tive site of dihydrolipoamide dehydrogenase in E 3 . Lipoamide is reoxidized in step 4 and
the regenerated coenzyme is repositioned in the active site of E x where it is ready to receive
a new two-carbon group. In these reactions, the lipoamide prosthetic group acts as a swing-
ing arm that visits the three active sites in the pyruvate dehydrogenase complex (Figure
13.2). The swinging arm portion of the E 2 subunit consists of a flexible polypeptide chain
that includes the lysine residue to which lipoamide is covalently bound.

The various subunits of the complex are arranged in a way that facilitates the
swinging arm mechanism of lipoamide. The mechanism ensures that the product of
one reaction does not diffuse into the medium but is immediately acted on by the next
component of the system. This is a form of channeling where the product of one reac-
tion becomes the substrate of a second reaction but it differs from other examples be-
cause, in this case, the two-carbon intermediate is covalently bound to the flexible
lipoamide group of E 2 .

The entire pyruvate dehydrogenase reaction is a series of coupled oxidation-reduction
reactions in which electrons are transported from the initial substrate (pyruvate) to the

oxidizing agent (NAD®). The four half reactions are

rot

acetyl CoA + C0 2 + H©

+ 2e° > pyruvate + CoA

-0.48

E 2 — lipoamide + 2H© +

2e® > E 2 — dihydrolipoamide

-0.29

E 3 — FAD + 2H© + 2e©

> e 3 — fadh 2

-0.34

NAD© + 2H© + 2e© —

NADH + H©

-0.32

(13.7)

Channeling and multienzyme complexes
were discussed in Section 5.11.

O Acetyl CoA

HS-CoA H 3 C — C — S-CoA

NADH + H®
NAD®

a Figure 13.2

Reactions of the pyruvate dehydrogenase complex. The lipoamide prosthetic group (blue) is attached by an amide linkage between lipoic acid and the
side chain of a lysine residue of E 2 . This prosthetic group is a swinging arm that carries the two-carbon unit from the pyruvate dehydrogenase active
site to the dihydrolipoamide acetyltransferase active site. The arm then carries hydrogen to the dihydrolipoamide dehydrogenase active site.

390

CHAPTER 13 The Citric Acid Cycle

▲ Figure 13.3

Structural model of the pyruvate dehydroge-
nase complex, (a) The inner core consists of
60 E 2 enzymes arranged in the shape of a
pentagonal dodecahedron with one E 2 trimer
at each of the 20 vertices. A single trimer is
outlined by a yellow box. The center of the
pentagon shape is indicated by the orange
pentagon. Note the linker regions projecting
upward from the surface of the core struc-
ture. (b) Cutaway view of the complete com-
plex showing the outer Ei enzymes (yellow)
and the BP-E 3 enzymes (red) located in the
space between the E 2 enzymes of the inner
core.

[From Zhou, H. Z. et al. (2001). The remarkable
structural and functional organization of the eu-
karyotic pyruvate dehydrogenase complexes.

Proc. Natl. Acad. Sci. (USA) 98:14082-14087.]

▲ A biochemistry laboratory.

Each half-reaction has a characteristic standard reduction potential (Table 10.4) that
provides some indication of the direction of electron flow. (Recall from Section 10.9
that the actual reduction potentials depend on the concentrations of reducing agents and
oxidizing agents.) Electron transport begins with pyruvate, which gives up two elec-
trons in the reverse of half- reaction 1. These electrons are taken up by E 2 -lipoamide.
Subsequent electron flow is from E 2 -lipoamide to E 3 -FAD to NAD®. The final product
is NADH, which carries a pair of electrons. There are many examples of metabolic pathway
enzymes with simple electron transport systems such as this one. They should not be
confused with the much more complex membrane-associated electron transport system
covered in the next chapter.

The pyruvate dehydrogenase complex is enormous. It is several times bigger than a
ribosome. In bacteria these complexes are located in the cytosol, and in eukaryotic cells
they are found in the mitochondrial matrix. Pyruvate dehydrogenase complexes are also
present in chloroplasts.

The eukaryotic pyruvate dehydrogenase complex is the largest multienzyme com-
plex known. The core of the complex is formed from 60 E 2 subunits arranged in the
shape of a pentagonal dodecahedron (12 pentagons joined at their edges to form a ball).
This shape has 20 vertices and each vertex is occupied by an E 2 trimer (Figure 13. 3 A).
Each of the E 2 subunits has a linker region projecting upward from the surface. This
linker contacts an outer ring of E : subunits that surround the inner core (Figure 13. 3B).
The linker region contains the lipoamide swinging arm.

The outer shell has 60 E x subunits. Each E x enzyme contacts one of the underlying
E 2 enzymes and makes additional contacts with its neighbors. The E x enzyme consists
of two a subunits and two /3 subunits (a 2 j8 2 ), so it is considerably larger than the E 2
enzyme of the core. The E 3 enzyme (an a 2 dimer) lies in the center of the pentagon
formed by the core E 2 enzymes. There are 12 E 3 enzymes in the complete complex, corre-
sponding to the 12 pentagons in the pentagonal dodecahedron shape. In eukaryotes, the
E 3 enzymes are associated with a small binding protein (BP) that’s part of the complex.

The model shown in Figure 13.3 has been constructed from high resolution elec-
tron microscopy images of pyruvate dehydrogenase complexes at low temperature
(cryo-EM) (Figure 13.1). In this technique, a large number of individual images
are combined and a three-dimensional image is built with the help of a computer.

Sample Calculation 13.1

Q. Calculate the standard Gibbs free energy change for the pyruvate dehydroge-
nase reaction.

A. From Equation 10.26, the overall change in standard reduction potential is

A cor a ror A ror

iaz: lal electron acceptor LAL electron donor

= -0.32 -(-0.48) = 0.16 V

from Equation 10.25,

AC°' = -nFAE°'

= -(2)(96.5)(0.16)

= -31 kj mol 1

The model is then matched with the structures of any of the individual subunits that
have been solved by X-ray crystallography or NMR. So far, it has not been possible to
grow large crystals of the entire pyruvate dehydrogenase complex on Earth and experi-
ments to grow crystals on the International Space Station in the absence of gravity were
also unsuccessful.

13.2 The Citric Acid Cycle Oxidizes Acetyl CoA 391

A similar pyruvate dehydrogenase complex is present in many species of bacteria
although some, such as gram- negative bacteria, have a smaller version where there are
only 24 E 2 enzymes in the core. In these bacteria, the core enzymes are arranged as a
cube with one trimer at each of the eight vertices. The E 2 subunits of the two different
bacterial enzymes and the eukaryotic mitochondrial and chloroplast versions are all
closely related. However, the gram-negative bacterial enzymes contain E x enzymes that
are unrelated to the eukaryotic versions.

Pyruvate dehydrogenase is a member of a family of multienzyme complexes known
as the 2-oxo acid dehydrogenase family. (Pyruvate is the smallest 2-oxo organic acid.)
We will encounter two other 2-oxo (or a-keto) acid dehydrogenases that closely resemble
pyruvate dehydrogenase in structure and function. One is a citric acid cycle enzyme,
a-ketoglutarate dehydrogenase (Section 13.3#4), and the other is branched chain
a-keto acid dehydrogenase, used in amino acid metabolism (Section 17.10E). All mem-
bers of the family catalyze essentially irreversible reactions in which an organic acid is
oxidized to C0 2 and a coenzyme A derivative is formed.

The reverse reactions are catalyzed in some bacteria by entirely different enzymes.
These reactions form part of a pathway for fixing carbon dioxide in anaerobic bacteria.
Some bacteria and some anaerobic eukaryotes convert pyruvate to acetyl CoA and C0 2
using pyruvate iferredoxin 2-oxidoreductase, an enzyme that is unrelated to pyruvate
dehydrogenase.

The regulation of pyruvate dehydroge-
nase is examined in Section 13.5.

KEY CONCEPT

Large multienzyme complexes improve
efficiency by channeling substrates and
products.

Acetyl CoA

S-CoA

Pyruvate + CoA + 2 Fd ox -> acetyl CoA + 2 Fd red + 2 H© (13.8)

HS-CoA

The terminal electron carrier in this case is reduced ferredoxin (Fd red ) and not NADH,
as with pyruvate dehydrogenase. The pyruvate iferredoxin oxidoreductase reaction is re-
versible and may be used to fix C0 2 by reductive carboxylation. Bacterial species that
have diverged very early in the history of life often contain pyruvate iferredoxin oxidore-
ductase and not pyruvate dehydrogenase suggesting that the former enzyme is more
primitive and pyruvate dehydrogenase evolved later.

13.2 The Citric Acid Cycle Oxidizes Acetyl CoA

Acetyl CoA formed from pyruvate or other compounds (such as fatty acids or some
amino acids) can be oxidized by the citric acid cycle. The eight reactions of the citric
acid cycle are listed in Table 13.1. Before examining each of the reactions individually,
we should consider two general features of the pathway; the flow of carbon and the pro-
duction of “high energy” molecules.

The fates of the carbon atoms are depicted in Figure 13.4. In the first reaction of the
citric acid cycle, the two -carbon acetyl group of acetyl CoA is transferred to the four-
carbon dicarboxylic acid oxaloacetate to form citrate, a six- carbon tricarboxylic acid.
The cycle proceeds with oxidative decarboxylation of a six- carbon acid and a five- carbon
acid. This releases two molecules of C0 2 and produces succinate, a four-carbon dicar-
boxylic acid. The remaining steps of the cycle convert succinate to oxaloacetate, the
original reactant that began the cycle.

The complete reactions are shown in Figure 13.5 where the two carbons of the
acetyl group are also colored green so their fate can be followed. Note that the two car-
bon atoms entering the cycle as the acetyl group on acetyl CoA are not the same carbon
atoms that are lost as C0 2 . However, the carbon balance in the overall reaction pathway
is such that for each two -carbon group from acetyl CoA that enters the cycle, two car-
bon atoms are released during one complete turn of the cycle. The two carbon atoms of
acetyl CoA become half of the symmetric four-carbon dicarboxylic acid (succinate) in
the fifth step of the cycle. The two halves of this symmetric molecule are chemically
equivalent so carbons arising from acetyl CoA become evenly distributed in molecules
formed from succinate.

Acetyl CoA is a “high energy” molecule (Section 10.8). The thioester linkage con-
serves some of the energy gained from the decarboxylation of pyruvate by the pyruvate
dehydrogenase complex. The net equation of the citric acid cycle (Table 13.1) tends to

Oxaloacetate

p i

? i

> or l

> c

QH 2 ^

^ NADH

^ • CO,

\ /

Plane of
symmetry

Succinate

GTP (or ATP)

NADH

• C0 2

▲ Figure 13.4

Fates of the carbon atoms from oxaloacetate
and acetyl CoA during one turn of the citric
acid cycle. The plane of symmetry of succi-
nate means that the two halves of the mole-
cule are chemically equivalent; thus, carbon
atoms from acetyl CoA (green) are uniformly
distributed in the four-carbon intermediates
leading to oxaloacetate. Carbon atoms from
acetyl CoA that enter in one turn of the cycle
are thus lost as C0 2 only in the second and
subsequent turns. Energy is conserved in the
reduced coenzymes NADH and QH 2 and in
one GTP (or ATP) produced by substrate level
phosphorylation.

392 CHAPTER 13 The Citric Acid Cycle

Table 13.1 The enzymatic reactions of the citric acid cycle

Reaction

Enzyme

1. Acetyl CoA + Oxaloacetate + H 2 0 » Citrate + HS-CoA + H©

Citrate synthase

2. Citrate Isocitrate

Aconitase (Aconitate hydratase)

3. Isocitrate + NAD© > a-Ketoglutarate + NADH + C0 2

Isocitrate dehydrogenase

4. a-Ketoglutarate + HS-CoA + NAD© > Succinyl CoA + NADH + C0 2

a-Ketoglutarate dehydrogenase complex

5. Succinyl CoA + GDP (or ADP) + P; Succinate + GTP(or ATP) + HS-CoA

Succinyl-CoA synthetase

6. Succinate + Q Fumarate + QH 2

Succinate dehydrogenase complex

7. Fumarate + H 2 0 L-Malate

Fumarase (Fumarate hydratase)

8. L-Malate + NAD© Oxaloacetate + NADH + H©

Malate dehydrogenase

Net equation:

Acetyl CoA + 3 NAD© + Q + GDP (or ADP) + P| + 2 H 2 Q > HS-CoA + 3 NADH + QH 2 + CTP (or ATP) + 2 C0 2 + 2 H©

obscure the fact that the citric acid cycle is equivalent to the oxidation of an acetyl CoA
molecule with release of electrons. The overall reaction sequence can be simplified to

S-CoA

1 o

C =0 + 2 H 2 0 + OH 0

ch 3

where the hydroxyl group is donated by inorganic phosphate in Reaction 5 and some of
the products are shown as free protons and free electrons. This form of the net equation
reveals that eight electrons are released during the oxidation. (Recall that oxidation re-
actions release electrons and reduction reactions take up electrons.) Six of the electrons
are transferred to three molecules of NAD® along with three of the protons depicted in
Reaction 13.9. The remaining two electrons are transferred to one molecule of
ubiquinone (Q) along with two of the protons. Two free protons are produced in each turn
of the cycle. (Keep in mind that the carbon dioxide molecules released during the citric
acid cycle do not actually come directly from acetyl CoA. Reaction 13.9 is a simplified
version that emphasizes the net oxidation.)

* 2 C0 2 + HS-CoA + 7 H 0 + 8e° (13.9)

BOX 13.2 WHERE DO THE ELECTRONS COME FROM?

Chemical reaction equations, such as Reaction 13.9, aren’t
very helpful in understanding where electrons are released
and taken up. In order to see the electron balance in such
reactions it’s often useful to redraw the structures with the
valence electrons replacing the lines that represent the chem-
ical bonds in most drawings. Each covalent bond involves
a shared pair of electrons and each of the standard atoms
(C, O, N, S) requires eight valence electrons. Covalently bonded
hydrogen atoms have only a single pair of electrons in their
single shell.

The oxidation of acetyl CoA from Equation 13.8 is
shown in this form in the figure. Note that only the electrons
in the outer shells of the atoms are shown. These are the ones
removed by oxidations or added in reduction reactions.
There are 42 electrons (21 pairs) in the reactants and 34 elec-
trons (17 pairs) in the products: C0 2 and Coenzyme A. Thus,

8 electrons are released in the oxidation. Most of the time,
electrons are released when double bonds are formed (as in

carbon dioxide) since this results in the sharing of an extra
electron pair.

CoA

C::0:

H:0:H

+ :p:H

H:C:H

H:0:H

18e 0

16e 0

<L>

0::C::0

" + H:S-CoA +7 H 0 + 8e 0
0::C::0

32e 0 2e 0 8e 0

▲ The oxidation of an acetyl CoA equivalent by
the citric acid cycle showing the valence elec-
trons in the reactants and products.

13.2 The Citric Acid Cycle Oxidizes Acetyl CoA 393

► Figure 13.5

Citric acid cycle. For each acetyl group that
enters the pathway, two molecules of C0 2 are
released, the mobile coenzymes NAD® and
ubiquinone (Q) are reduced, one molecule of
GDP (or ADP) is phosphorylated, and the ac-
ceptor molecule (oxaloacetate) is re-formed.

Oxidation

NADH + H

NAD

Hydration

Oxidation

Malate

dehydrogenase

COO^

HO— C — H

ch 2

coo'

L-Malate

H 2 0

coo

H — C
C — H

Fumarase

,©

COO'

Fumarate

QH 2

FAD

Succinate

dehydrogenase

complex

COO'

ChH,

CH 2

COO'

,©

Succinate

HS-CoA

Substrate-level GTP (or ATP)
phosphorylation GDp (or ADP)

Succinyl-CoA

synthetase

Oxaloacetate

coo°

ChH,

ch 2

C = 0

Entry of substrate
by condensation
with oxaloacetate

COO^

Citrate

Aconitase

COO'"

CH 2

H — C — COO 0

HO— C — H

COO'
Isocitrate

(D V - NAD@

Isocitrate naqh
dehydrogenase

^ co 2

coo 0

CH 2

ch 2

C = 0

a-Ketoglutarate
dehydrogenase
complex

COO'
a-Ketoglutarate

Rearrangement

First

oxidative

decarboxylation

HS-CoA

NAD 0

Second

oxidative

NADH

decarboxylation

C0 2

S-CoA
Succinyl CoA

394 CHAPTER 13 The Citric Acid Cycle

KEY CONCEPT

The citric acid cycle is a mechanism for
the oxidation of the acetyl group of
acetyl CoA.

Most of the energy released in the citric acid cycle reactions is conserved in the
form of electrons transferred from organic acids to generate the reduced coenzymes
NADH and QH 2 (Figure 13.5). NADH is formed by the reduction of NAD® at three
oxidation-reduction steps — two of these are oxidative decarboxylations. QH 2 is formed
when succinate is oxidized to fumarate. Subsequent oxidation of the reduced coen-
zymes by membrane-associated electron transport leads to the transfer of electrons
from NADH and QH 2 to a terminal electron acceptor. In the case of most eukaryotes
(and many prokaryotes), this terminal electron acceptor is oxygen, which is reduced to
water. Membrane-associated electron transport is coupled to the production of ATP
from ADP and Pj. The entire process (electron transport + phosphorylation of ADP) is
often referred to as oxidative phosphorylation when oxygen is present (Chapter 14). In
addition to the formation of reducing equivalents, the citric acid cycle produces a
nucleotide triphosphate directly by substrate level phosphorylation. The product can be
either ATP or GTP, depending on the cell type or species.

13.3 The Citric Acid Cycle Enzymes

The citric acid cycle can be viewed as a multistep catalytic reaction returning to its orig-
inal state after an acetyl CoA molecule is oxidized. This view is based on the fact that
when the reactions operate as a cycle the original reactant, oxaloacetate, is regenerated.
By definition, a catalyst increases the rate of a reaction without itself undergoing net
transformation. All enzymatic reactions, in fact all catalytic reactions, can be repre-
sented as cycles. An enzyme goes through a cyclic series of conversions and finally
returns to the form in which it began. In this sense, the citric acid cycle fits the description
of a catalyst.

Taken as a whole, the citric acid cycle is a mechanism for oxidizing the acetyl group
of acetyl CoA to C0 2 by NAD® and ubiquinone. When the citric acid cycle operates
in isolation its intermediates are re-formed with each full turn of the cycle. As a result,
the citric acid cycle doesn’t appear to be a pathway for net synthesis or degradation of
any of the intermediates in the pathway unlike, for example, the gluconeogenesis path-
way or the glycolysis pathway. However, we will see later on (Section 13.6) that the citric
acid pathway doesn’t always operate in isolation and appearances can be deceiving.
Some of the intermediates are shared with other pathways. Let’s first examine the cat-
alytic aspect of the citric acid cycle by examining each of the eight enzymatic steps.

v Figure 13.6

Reaction catalyzed by citrate synthase. In the

first step, acetyl CoA combines with
oxaloacetate to form an enzyme-bound
intermediate, citryl CoA. The thioester is
hydrolyzed to release the products, citrate
and HS-CoA.

1. Citrate Synthase

In the first reaction of the citric acid cycle, acetyl CoA reacts with oxaloacetate and
water to form citrate, HS-CoA, and a proton. This reaction is catalyzed by citrate syn-
thase and results in the formation of an enzyme-bound intermediate called citryl CoA
(Figure 13.6).

Citrate is the first of two tricarboxylic acids that are part of the cycle. The standard
Gibbs free energy change for the citrate synthase reaction is —31.5 kj-mol -1 (AG°' = — 31.5
kj-mol -1 ) due to the hydrolysis of the high energy thioester bond in the citryl CoA inter-
mediate. Normally you might expect that such a large negative Gibbs free energy change

coo®

S-CoA

C = 0

ch 2 +

coo®

ch 3

Acetyl Cc

Oxaloacetate

S-CoA

c = o
I

r 2

HO — C — COO'

ch 2

coo°

Citryl CoA

,©

COO G

h 2 o ch 2

^ > HO — c— COO® + HS-CoA + H©

ch 2

coo®

Citrate

13.3 The Citric Acid Cycle Enzymes 395

would be coupled to synthesis of ATP — keeping in mind that the actual Gibbs free energy
change inside the cell might be very different. Indeed, the hydrolysis of the similar
thioester bond in succinyl CoA (Reaction 5 of the citric acid cycle) is coupled to synthesis
of GTP (or ATP). However, in the case of the citrate synthase reaction, the available energy
is used for a different purpose. It ensures that the reaction proceeds in the direction of cit-
rate synthesis when the concentration of oxaloacetate is very low (Figure 13.7). This ap-
pears to be the normal situation when the citric acid cycle is operating. In the presence of
only small (catalytic) amounts of oxaloacetate the equilibrium of the reaction depicted in
Figure 13.6 still favors citrate synthesis. In other words, the actual Gibbs free energy
change inside the cell is close to zero. The reaction is a near-equilibrium reaction. The
thermodynamics ensures that the citric acid cycle operates in the direction of acetyl CoA
oxidation even under conditions where the concentration of oxaloacetate is very low.

Citrate synthase is a transferase — one of the six categories of enzymes described in
Section 5.1. Transferases catalyze transfer reactions, in this case transfer of an acetyl
group. The term “synthase” is used for transferases that do not use ATP as a cofactor.
“Synthetases,” on the other hand, are members of the ligase category of enzymes
(Section 5.1). The reactions catalyzed by synthetases must be coupled to ATP (or GTP)
hydrolysis. It’s important to remember the difference between synthases and synthetases
since the words look very similar and since the citric acid cycle contains an example of
each type of enzyme. (For some reason, it’s easier to pronounce “synthetase” and it’s
tempting to throw in the extra syllable when you should be saying “synthase .”)

In gram-positive bacteria, archaebacteria, and eukaryotes, citrate synthase is a
dimeric protein composed of two identical subunits. In gram-negative bacteria, the
enzymes are hexameric complexes of identical subunits.

In animals each subunit of the enzyme has two distinct domains: a small flexible
domain on the outer surface and a larger domain that forms the core of the protein
(Figure 13.8). The two subunits associate by interactions between four a helices in each
of the large domains to form an a helix sandwich. Citrate synthase undergoes a large
conformational change on binding oxaloacetate as shown in Figure 13.8. The binding
site lies at the base of a deep cleft between the small domain of one subunit and the
large domain of the other subunit. When oxaloacetate is bound, the small domain ro-
tates by 20° relative to the large domain. This closure creates the binding site for acetyl
CoA — a site which is formed by amino acid side chains from both large and small
domains. When the reaction is complete coenzyme A is released. The enzyme then
reverts to the open conformation when citrate is released.

The structure of the enzyme requires that oxaloacetate and acetyl CoA bind sequen-
tially. This reduces the chance of binding acetyl CoA in the absence of oxaloacetate and

(a)

Small domain

(b)

° + o

Oxaloacetate Acetyl CoA

+ o

20° rotation

HS-CoA FT

▲ Figure 13.7

Representation of the relative ratios of products
and reactants in the citrate synthase reaction.

The equilibrium constant (/C eq ) for the cit-
rate synthase reaction can be calculated
from standard Gibbs free energy change ac-
cording to Equation 1.12, K eq = 2.7 x 10 5 ,
meaning that, at equilibrium, the concentra-
tions of products are more than 200,000
times that of the reactants. [Not to scale.]

v Figure 13.8

Citrate synthase induced fit mechanism. The

two identical subunits are colored blue and
purple. Each is composed of a small and a
large domain, (a) Open conformation. The
substrate binding site is located in the deep
cleft between the small domain of one sub-
unit and the large domain of the other.

[PDB 5CSC] (b) Closed conformation. The
small domain has shifted relative to the
large domain in order to close off the large
binding cleft seen in the open conformation.
Substrate analogues are shown as space-fill-
ing models. This version of the enzyme is
from chicken {Gallus gallus). [PDB 6CSC]

Large domain

396 CHAPTER 13 The Citric Acid Cycle

BOX 13.3 CITRIC ACID

The discovery of citric acid is usually attributed to Abu Musa
Jabir ibn Hayyan (-721 — 815), known as Geber in Europe. He
worked in Kufa in modern-day Iraq and is recognized as the
father of modern chemistry. Jabir identified citric acid as a
major component of citrus fruits such as lemons and limes. We
know now that the level of citric acid in these fruits is related to
its ability to act as a preservative and a reservoir of carbon. This
is unrelated to the role of citrate in the citric acid cycle.

Citric acid is a weak organic acid (pJ^ a i = 3.2, pi^ a 2 = 4.8,
pIC a3 = 6.4). The sodium salt is sometimes used as a buffer in
biochemistry labs and in drugs but its most important appli-
cation is as a food additive, especially in soft drinks.

▲ Citric acid is an important natural preservative in citrus fruits.

the possibility of catalyzing hydrolysis of the thioester bond of acetyl CoA in a wasteful
reaction. This potential side reaction is a very real danger since the thioester bond of
acetyl CoA is near the active site for hydrolysis of the citryl CoA thioester and since the
concentration of oxaloacetate may be very low relative to that of acetyl CoA. Our previ-
ous examples of an induced fit mechanism involved protecting ATP from inappropriate
hydrolysis but the same principle applies here. We will encounter several other examples
of important structure-function relationships in this chapter and the next one.

2. Aconitase

KEY CONCEPT

Stereospecific reactions occur because
substrates bind to enzymes in specific
orientations.

Aconitase (systematic name: aconitate hydratase) catalyzes a near- equilibrium conversion
of citrate to isocitrate. Citrate is a tertiary alcohol and thus cannot be oxidized directly
to a keto acid. The formation of a keto acid intermediate is required for the oxidative
decarboxylation reaction that occurs in step 3 of the citric acid cycle. The step catalyzed
by aconitase creates a secondary alcohol in preparation for step 3. The name of the enzyme
is derived from ds-aconitate, an enzyme-bound intermediate of the reaction. The reac-
tion proceeds by the elimination of water from citrate to form a carbon-carbon double
bond. This is followed by stereospecifc addition of water to form isocitrate.

coo°

▲ Figure 13.9

Structure of 2R,3S-isocitrate.

ch 2

HO — C— COO°

COO 0

Citrate

H 2 0

h 2 o

C — COO°

/ \ n

coo°

c/s- Aconitate

H 2 0 ch 2

^ HC— COO°

~T~ HO-CH (13-10)

H2 ° 1 O

COO 0

Isocitrate

The aconitase gene is a member of a complex gene family. The family encodes dis-
tinct mitochondrial and cytoplasmic versions of aconitase, a regulatory protein with no
catalytic activity, and an enzyme involved in the synthesis of amino acids (Sections 13.8
and 17.3C). Bacteria contain two distantly related enzymes, aconitase A and aconitase B.
All family members contain a characteristic [4 Fe-4 S] iron-sulfur cluster. In the next
chapter we will encounter many oxidation-reduction enzymes with iron-sulfur clus-
ters. In most of these oxidation-reduction enzymes, the iron-sulfur clusters participate
in electron transport but members of the aconitase family are unusual because the role
of the iron-sulfur cluster is to aid in the binding of citrate. The aconitase reaction is an
isomerization reaction and not an oxidation- reduction reaction.

Note that citrate is not a chiral molecule because none of the carbon atoms is
bonded to four different groups. However, the product of the reaction, isocitrate, has two
chiral centers, C2 and C3. Each of these carbon atoms has four different constituents

13.3 The Citric Acid Cycle Enzymes 397

BOX 13.4 THREE POINT ATTACHMENT OF PROCHIRAL SUBSTRATES TO ENZYMES

When the citric acid cycle was first proposed by Krebs, the
inclusion of the citrate-to- isocitrate reaction was a major
barrier to its acceptance because labeling studies indicated
that only one of the two possible forms of 2R,3S-isocitrate
was produced in cells. The “problem” was not that a chiral
molecule was produced from a non- chiral molecule — this is
easily understood. The difficulty was in understanding why
formation of the double bond of ds-aconitate, and subse-
quent addition of water to form isocitrate, occurred only in
the moiety contributed originally by oxaloacetate and not in
the group derived from acetyl CoA. When isotopically
labeled acetate was added to cells the 14 C-labeled carbon atoms
appeared in citrate as shown in green in Reaction 13.10.
Since citrate is a symmetric molecule, the labeled carbon
atoms were expected to show up equally in the two versions
of isocitrate shown in the figure on the right.

Instead, only the left-hand form was produced. At the
time, conversion of a non- chiral molecule to a single form of
chiral isomer was unknown but in 1948, Alexander Ogston
showed how the active site of an enzyme could distinguish
between chemically equivalent groups on the citrate mole-
cule. Ogston envisioned citrate binding in a manner he called
three point attachment, with nonidentical groups involved in
the enzyme-substrate binding (see figure). Once citrate
is correctly bound to the asymmetric binding site, the
two — CH 2 — COO® groups of citrate have specific orienta-
tions and thus are no longer equivalent. Formation of the
carbon-carbon double bond can only take place in the group
contributed by oxaloacetate.

coo 0 coo 0

I I

ch 2 ch 2

HC — COO 0 HC — COO 0

I I

HO — CH HO — CH

COO 0 COO 0

▲ Two forms of isocitrate. The green carbon atoms represent the group
originally derived from acetyl CoA. The reaction catalyzed by aconi-
tase was expected to yield two forms of isocitrate in equal quantities
because the substrate (citrate) is symmetric. Only the left-hand form
was produced.

Citrate is a prochiral molecule because it can react asym-
metrically in spite of the fact that it is chemically symmetric.
There are now many examples of such reactions in metabolic
pathways.

reactive group

▲ Binding of citrate to the active site of aconitase. The central carbon
atom of the citrate molecule is shown with four attached groups: the
hydroxyl group ( — OH) is represented by a square; the carboxyl group
( — COOH) by a triangle; the two — CH2 — COO — groups are shown as
spheres. The two — CH2 — COO — groups are chemically indistinguish-
able, but the one derived from acetyl CoA is shown as a green sphere
and the one derived from oxaolacetate is colored blue. A cartoon of
aconitase is depicted as an asymmetric molecule with three-point at-
tachments sites for the hydroxyl group, the carboxyl group, and one
of the — CH2 — COO — groups. When citrate is oriented as shown in
the top figure, it can bind to aconitase and the reaction takes place
in the moiety derived from oxaloacetate. The other orientation (bottom)
cannot bind to the enzyme and the reaction cannot take place in the
group derived from acetyl CoA.

and in each case the four groups can be arranged in two different orientations. There
are four different stereoisomers of isocitrate but only one of them is produced in the re-
action catalyzed by aconitase. The formal name of this product is 2R,3S-isocitrate
(Figure 13.9) using the RS nomenclature described in Box 3.2. This is one of the few
times when this nomenclature is useful in introductory biochemistry.

3. Isocitrate Dehydrogenase

Isocitrate dehydrogenase catalyzes the oxidative decarboxylation of isocitrate to form The regulation of isocitrate dehydroge-
a-ketoglutarate (Figure 13.10). This reaction is the first of four oxidation-reduction nase in prokaryotes is described in

reactions in the citric acid cycle. The reaction is coupled to the reduction of NAD® and Section 1 3.8.

occurs in two steps involving an enzyme-bound oxalo succinate intermediate.

398 CHAPTER 13 The Citric Acid Cycle

COO G

<=H 2

HC — COO 0 + NAD®

HO — CH

COO©

Isocitrate

dehydrogenase

coo®

ch 2

HC-

✓

o©

c=o
coo©

Oxalosuccinate

+ H© + NADH

H ©

COO©

ch 2 + co 2

C =0

coo©

a-Ketoglutarate
▲ Figure 13.10

Isocitrate dehydrogenase reaction. The enzyme
catalyzes an oxidation-reduction reaction
using NAD© as the electron acceptor.
Oxalosuccinate is an unstable intermediate
that is rapidly decarboxylated to C0 2 and
a-ketoglutarate. This is the first decarboxy-
lation step in the citric acid cycle.

KEY CONCEPT

The important “pay off” reactions of the
citric acid cycle are those that produce
reducing equivalents such as NADH
and QH 2 .

In the first step, the alcohol group of isocitrate is oxidized by removal of two hydro-
gens to form a — C = O double bond. This is a typical dehydrogenase reaction. One of
the hydrogens (the one bound to the carbon atom) is transferred to NAD® as a hydride
ion carrying two electrons and the other (the one on the — OH group) is incorporated
into the final product. This is the first of the reactions that result in the loss of electrons
(i.e., oxidation of an organic acid).

Oxalosuccinate, an unstable keto acid, is the product of the first step in the overall
reaction catalyzed by a-ketoglutarate dehydrogenase. Before it is released from the
enzyme, the intermediate undergoes decarboxylation to form a-ketoglutarate in the
second step of the reaction. The decarboxylation reaction is associated with the release
of C0 2 and uptake of a proton. The overall stoichiometry of the reaction is

Isocitrate + NAD® > a-Ketoglutarate + NADH + C0 2 (13.11)

There are several different versions of isocitrate dehydrogenase. Bacteria contain
both an NAD® -dependent enzyme and an NADP® -dependent enzyme. Eukaryotes
also have both types but, in addition, the NADP® -dependent enzymes form several
subclasses. In general, the NAD® -dependent enzyme is localized to the mitochondria
and plays the major role in the citric acid cycle. The NADP® -dependent enzymes are
found in the cytoplasm, chloroplasts, and other membrane compartments. All forms of
the enzymes are homologous by sequence similarity and they share a common ancestor
with an enzyme in the leucine biosynthesis pathway (Section 13.9, Section 17.3C).

4. The a-Ketoglutarate Dehydrogenase Complex

Oxidative decarboxylation of a-ketoglutarate is analogous to the reaction catalyzed by
pyruvate dehydrogenase. In both cases, the reactants are an a-keto acid and HS-CoA
and the products are C0 2 and a “high energy” thioester compound. Step 4 of the citric
acid cycle is catalyzed by a-ketoglutarate dehydrogenase (also known as 2-oxoglutarate
dehydrogenase) (Figure 13.11)

a-Ketoglutarate dehydrogenase is a large complex that resembles pyruvate dehy-
drogenase in both structure and function. The same coenzymes are involved and the
reaction mechanism is the same. The three component enzymes of the a-ketoglutarate
dehydrogenase complex are a-ketoglutarate dehydrogenase (E 1? containing TDP), di-
hydrolipoamide succinyl transferase (E 2 , containing a lipoamide swinging arm), and
dihydrolipoamide dehydrogenase (E 3 , the same flavoprotein found in the pyruvate
dehydrogenase complex). The overall reaction is the second of the two C0 2 producing
reactions in the citric acid cycle and the second reaction that generates reducing equiva-
lents. In the four remaining reactions of the cycle, the four-carbon succinyl group of
succinyl CoA is converted back to oxaloacetate.

Eukaryotic cells have a single mitochondrial a-ketoglutarate dehydrogenase.
Archaebacteria, and some other species of bacteria, do not have a-ketoglutarate dehy-
drogenase. Instead, they convert a-ketoglutarate to succinyl CoA using an entirely
different enzyme called 2-oxoglutarate:ferredoxin oxidoreductase.

5. Succinyl CoA Synthetase

The conversion of succinyl CoA to succinate is catalyzed by succinyl CoA synthetase,
sometimes called succinate thiokinase. The reaction couples hydrolysis of the thioester
linkage in succinyl CoA to formation of a nucleoside triphosphate — either GTP or ATP,
depending on the species. The complicated IUPAC names of these two related enzymes
are: succinate-CoA ligase, ADP- forming (E.C. 6.2. 1.5); and succinate-CoA ligase, GDP-
forming (E.C. 6.2. 1.4).

Inorganic phosphate is one of the reactants and the reaction takes place in three
steps (Figure 13.12).

The first step generates succinyl phosphate as an intermediate and releases coen-
zyme A. In the second step, the phosphoryl group is transferred to a histidine side chain
in the active site of the enzyme to form a stable phosphoenzyme intermediate. The third
step transfers the phosphoryl group to GDP to form GTP. This reaction is the only
example of substrate level phosphorylation in the citric acid cycle. (Recall from Section 10.8

13.3 The Citric Acid Cycle Enzymes 399

that the standard Gibbs free energy change for hydrolysis of the thioester linkage in
succinyl CoA is approximately equivalent to that of ATP hydrolysis.) The overall stoi-
chiometry of the succinyl CoA synthetase reaction is

Succinyl CoA + Pj + GDP > Succinate + HS-CoA + GTP (13.12)

Inorganic phosphate contributes the phosphoryl group to GDP, plus an oxygen to form
succinate and a hydrogen to form HS-CoA. Note that the enzyme is named for the re-
verse reaction where succinyl CoA is synthesized from succinate at the expense of GTP
or ATP. It is called a synthetase because the reaction combines two molecules and it is
coupled to hydrolysis of nucleoside triphosphate.

The enzyme is composed of two a and two /? subunits (Gk/fe)- The /? subunits con-
tain the binding site for the nucleotide. Bacterial versions use ATP while animals often
have two versions of the enzyme — one that uses GTP and one that uses ATP. They dif-
fer in their /? subunits. The GTP- dependent versions clearly have evolved from the ATP-
dependent versions. It’s not clear why animal mitochondria have two versions of suc-
cinyl CoA synthetase in their mitochondria but one possibility is that the ATP-dependent
version is used in the citric acid cycle and the GTP-dependent version primarily cat-
alyzes the reverse reaction in some cells. Archaebacteria, and some other bacteria, do
not have succinyl CoA synthetase. They carry out a similar reaction using an entirely
different enzyme.

6. Succinate Dehydrogenase Complex

Succinate dehydrogenase catalyzes the oxidation of succinate to fumarate forming a
carbon-carbon double bond with the loss of two protons and two electrons (Figure 13.13).
The protons and electrons are passed to a quinone, which is reduced to QH 2 . (Ubiquinone
is the preferred substrate in almost all cases but some bacteria use menaquinone.) The
enzyme is present in all species and FAD is an essential bound cofactor.

One important feature of this reaction is the scrambling of the original acetyl car-
bon atoms. They can no longer be specifically identified (i.e., green) in the symmetrical

COO G

r 2

CH 2 + HS-CoA + NAD®
C =0

coo®

u-Ketoglutarate

\ /

coo®

ChH 2

CH 2 + C0 2 + NADH

c=o

S-CoA
Succinyl CoA

▲ Figure 13.11

Reaction catalyzed by a-ketoglutarate dehy-
drogenase. This is similar to the reaction
catalyzed by pyruvate dehydrogenase.

The structure of menaquinone is shown
in Figure 14.21.

BOX 13.5 WHAT’S IN A NAME?

a-Ketoglutarate is clearly named after the five -carbon dicar-
boxylic acid glutarate ( e OOC— CH 2 — CH 2 — CH 2 — COO 0 ).
The keto group is on the a carbon or the first carbon after
one of the carboxyl groups. This naming convention is sim-
ilar to the one we encountered in naming ct-amino acids
(Section 3.1). As is the case with amino acids, the correct
chemical name, or systematic name, for a-ketoglutarate
could be cc 2 -keto glutarate.” However, the formal name is ac-
tually 2 -oxo glutarate since according to the IUPAC/IUBMB
rules of nomenclature the term “keto” should now be avoided.

It is perfectly acceptable to refer to organic molecules by
their common (trivial) names if these common names are
well known. For example, if you look back to step 1 of the
citric acid cycle you can see that the systematic name for
oxaloacetate is 2-oxosuccinate since it is a derivative of the
four-carbon dicarboxylic acid, succinate. “Oxaloacetate” is
the well-known and accepted common name for this com-
pound and it would be confusing to use any other name.
When it comes to the correct name for a-ketoglutarate, the
situation is more complicated because a-ketoglutarate is the
old-fashioned systematic name of the molecule and the new

rules say that the systematic name should be 2 -oxo glutarate.
The new name is becoming more and more popular in the
scientific literature. Here, we continue to use the well-known
name a-ketoglutarate on the grounds that it has become an
acceptable common name for this compound. It’s very likely
that this will change in future editions.

400 CHAPTER 13 The Citric Acid Cycle

◄ Figure 13.12

Proposed mechanism of succinyl CoA syn-
thetase. Phosphate displaces CoA from a
bound succinyl CoA molecule, forming the
mixed acid anhydride succinyl phosphate
as an intermediate. The phosphoryl group
is then transferred from succinyl phosphate
to a histidine residue of the enzyme to form
a relatively stable covalent phosphoenzyme
intermediate. Succinate is released, and the
phosphoenzyme intermediate transfers its
phosphoryl group to GDP (or ADP, depending
on the organism), forming the nucleoside
triphosphate product.

coo°

H©

▲ GTP-dependent succinyl CoA synthetase. The

structure of one unit of the dimer is shown
with the a and ft subunits in different colors.
A molecule of GTP is bound at the active
site within the ft subunit. This is the pig
{Sus scrofa ) version of the enzyme.

[PDB 2FPG]

reactant, succinate, or in the product, fumarate. This has interesting consequences (see
Problem #6).

The active site of the enzyme is formed from two different subunits. One subunit
contains iron-sulfur clusters and the other is a flavoprotein with covalently bound FAD.
The succinate dehydrogenase dimer is bound to two membrane polypeptides to form a
larger complex. The membrane components consist of a cytochrome b , with its associ-
ated heme group, and a quinone binding site. The electron transport cofactors partici-
pate in the transfer of electrons from succinate to FAD to several iron-sulfur clusters to
heme to the quinone.

Recall that FADH 2 in subunit E 3 of pyruvate dehydrogenase is reoxidized by NAD 0
to complete the catalytic cycle of that enzyme. In the succinate dehydrogenase reaction,
FADH 2 is reoxidized by Q to regenerate FAD. In the past, it was very common to show
FADH 2 as the redox product of this reaction but since FAD is covalently bound to the
enzyme, the catalytic cycle is not completed until bound FADH 2 is reoxidized and the
mobile product QH 2 is released.

The succinate dehydrogenase reaction is unusual for a dehydrogenase because it
uses ubiquinone as an electron acceptor (oxidizing agent) instead of NAD®. It is also
unusual in many other ways, as we will see in the next chapter. The succinate dehydroge-
nase complex is part of the electron transport system located in the plasma membrane of
prokaryotes and in the inner mitochondrial membrane in eukaryotic cells. We will discuss
this enzyme in more detail in Section 14.6 and examine its structure (Figure 14.9). In
bacteria, the bulk of the enzyme complex projects into the cytoplasm where it can bind
succinate and release fumarate as part of the citric acid cycle. In mitochondria, the
active site is on the matrix side of the membrane where the other citric acid cycle en-
zymes are located.

13.3 The Citric Acid Cycle Enzymes 401

The substrate analog malonate is a competitive inhibitor of the succinate dehydro-
genase complex as described in Section 5. 7 A. Malonate, like succinate, is a dicarboxylate
that binds to cationic amino acid residues in the active site of the succinate dehydroge-
nase complex. However, malonate cannot undergo oxidation because it lacks the
— CH 2 — CH 2 — group necessary for dehydrogenation. In experiments with isolated
mitochondria or cell homogenates, the presence of malonate caused succinate, a-
ketoglutarate, and citrate to accumulate. Such experiments provided some of the orig-
inal evidence for the sequence of reactions in the citric acid cycle.

7. Fumarase

Fumarase (systematic name: fumarate hydratase) catalyzes the near-equilibrium con-
version of fumarate to malate through the stereospecific trans addition of water to the
double bond of fumarate.

H COO°

\ /

c. Fumarase

II + h 2 o <

°OOC H

.coo

©,

000

OH
C"

-C..,

i ""'H

(13.13)

Fumarate L-Malate

Fumarate is a prochiral molecule. When fumarate is positioned in the active site of
fumarase, the double bond of the substrate can be attacked from only one direction. The
product of the reaction is exclusively the L stereoisomer of the hydroxy acid malate.

There are two unrelated fumarases that can catalyze the same reaction. The class I
enzyme is found in most bacteria. The class II enzyme is present in some bacteria and
all eukaryotes. Some bacteria, such as E. coli , have both forms of the enzyme. One form
is active in the normal citric acid cycle pathway and the other usually specializes in the
reverse reaction to convert malate to fumarate.

8. Malate Deydrogenase

The last step in the citric acid cycle is the oxidation of malate to regenerate oxaloacetate,
with formation of a molecule of NADH.

coo°

ch 2

1 ©

coo®

Succinate

+ Q

Succinate

dehydrogenase

H COO'

I©

+ QH 2

©,

ooc

Fumarate

▲ Figure 13.13

The succinate dehydrogenase reaction.

a Green (unripe) apples. The sour taste of un-
ripe apples is mostly due to the presence
of malate. Malic acid was first isolated from
apple juice and it was named after the Latin
word for apple {malum).

HO — C — H Malate C=0

dehydrogenase I

CH 2 + NAD® ; t CH 2 + NADH + H®

COO® coo®

L-Malate Oxaloacetate

(13.14)

BOX 13.6 ON THE ACCURACY OF THE WORLD WIDE WEB

There’s lots of good stuff on the web but everyone should be cautious about the
quality of some webpages. The citric acid cycle is a fun test case for accuracy. Most
sites get the basics correct but students are often challenged to find a website that
accurately depicts every reaction of the pathway with no errors — including balanc-
ing every equation. Can you find such a website? The most common errors are
leaving out protons and QH 2 .

The one site you can rely on is the IUBMB Enzyme Nomenclature site that lists
the correct reactions for each enzyme in the citric acid cycle: www.chem.qmul.ac.uk/
iubmb/enzyme/

Some instructors have been known to give extra marks to students who can find a com-
pletely accurate website. Some students have been known to create their own webpages.

We consider the transfer of reducing
equivalents to Q again in Chapter 14,
where we will see the role of the suc-
cinate dehydrogenase complex in
membrane-associated electron transport.

The evolutionary origin of fumarase
and the significance of the reverse
reaction in bacteria are described in
Section 13.8.

402

CHAPTER 13 The Citric Acid Cycle

This reaction is catalyzed by NAD® -dependent malate dehydrogenase. The near-equi-
librium interconversion of the a-hydroxy acid L-malate and the keto acid oxaloacetate is
analogous to the reversible reaction catalyzed by lactate dehydrogenase (Sections 7.3
and 1 1.3B). This is not surprising since lactate dehydrogenase and malate dehydrogenase
are homologous — they share a common ancestor.

The standard Gibbs free energy change for this reaction is +30 kj mol -1 (AG°' =
30 kj mol -1 ). Since this is a near-equilibrium reaction it means that under the condi-
tions found inside the cell, the concentration of malate is very much higher than that of
oxaloacetate. We’ve seen in the case of the citrate synthase reaction that the low concen-
The structures of malate dehydrogenase tration of oxaloacetate explains the Gibbs free energy change of that reaction. In the

and lactate dehydrogenase are com- next section we’ll see how the low concentration of oxaloacetate relative to that of

pared in Figure 4.22. malate explains some transport pathways.

13.4 Entry of Pyruvate Into Mitochondria

In bacterial cells, pyruvate is converted to acetyl CoA in the cytosol but in eukaryotic
cells the pyruvate dehydrogenase complex is located in mitochondria (and in chloro-
plasts). Since glycolysis takes place in the cytoplasm, pyruvate must first be imported
into the mitochondria (or chloroplasts) so that it can serve as a substrate in the reac-
tion. The mitochondrion is enclosed by a double membrane. Small molecules such as

BOX 13.7 CONVERTING ONE ENZYME INTO ANOTHER

Despite having a low sequence identity, lactate dehydrogenase and malate dehydroge-
nase are closely related in three-dimensional structure and they clearly have evolved
from a common ancestor. These enzymes catalyze reversible oxidation of 2 -hydroxy
acids that differ by only one carbon (malate has an additional carboxylate attached to
C-3 of lactate). Both enzymes are highly specific for their own substrates. However,
site specific mutation of a single amino acid residue of the lactate dehydrogenase of
Bacillus stearothermophilus changes this enzyme to a malate dehydrogenase (see
figure). Conversion of Gin- 102 to Arg-102 completely reverses the specificity of the
dehydrogenase. The positively charged side chain of the arginine forms an ion pair
with the 4-carboxylate group of malate, and the mutant enzyme becomes inactive
with lactate.

coo®

l 2

HO — C — H

ch 3

L-Lactate

NAD® NADH, H® COO®

C =0

Lactate dehydrogenase

ch 3

Pyruvate

COO G NAD® NADH, H® COO G

I W I

HO — C — H a C=0

Malate dehydrogenase

CH,

COO

L-Malate

>©

ch 2

coo°

Oxaloacetate

▲ Orientation of the substrate molecule in the active site of lactate dehydrogenase from Bacillus stear-
mothermophilus. (a) The three-carbon substrate pyruvate bound to the native enzyme. Neither ox-
aloacetate nor malate can bind at this site, (b) The four-carbon substrate oxaloacetate bound to
the Gln-to-Arg mutant (position 102).

(a) — Gln-102—

c — nh 2

ch 3

1 H v=

O — C Xmdh

I H

0-4-0

H 2 N, ©,, nh 2

— Arg-171—

(b) — Arg-102-

H 2 N'"© nh 2
oX?+o
I

ch 2

1 H v=

0 = C Xnadh

I H

o4o

h 2 n^©^ nh 2

-Arg-171 —

13.4 Entry of Pyruvate Into Mitochondria 403

◄ Figure 13.14

Import of pyruvate and export of PEP. Pyruvate
is imported into mitochondria from the cyto-
plasm via a pyruvate transporter located in
the inner mitochondrial membrane. Phos-
phoenolpyruvate (PEP) is exported to the
cytoplasm via a PEP transporter.

pyruvate pass through the outer membrane via aqueous channels formed by trans-
membrane proteins called porins (Section 9.11A). These channels allow free diffusion
of molecules with molecular weights less than 10,000. However, in order to pass
through the inner membrane a specific transport protein is required for most metabo-
lites. Pyruvate translocase specifically transports pyruvate in symport with H®. Once
inside the mitochondrion, pyruvate can be converted to acetyl CoA and C0 2 . In
eukaryotic cells the enzymes of the citric acid cycle are also located in the mitochondria
(Figure 13.14).

Recall that one of the intermediates in the citric acid cycle is oxaloacetate and it can
also be a substrate for gluconeogenesis. Since gluconeogenesis is a cytoplasmic pathway,
it’s necessary to move oxaloacetate, or its equivalent, from the mitochondria to the
cytoplasm. In mammals this is accomplished using a mitochondrial version of phos-
phoenolpyruvate carboxykinase (PEPCK), that converts oxaloacetate to phospho-
enolpyruvate (PEP). Mitochondria possess a PEP transporter that moves PEP to the
cytoplasm (Figure 13.14). It would be very inefficient to transport oxaloacetate directly
because its concentration in the mitochondria is very low compared to its concentration
in the cytoplasm. (Deficiencies in the human mitochondrial PEPCK lead to death
within the first two years of life.)

There are two other problems associated with the compartmentation of the
citric acid cycle in mitochondria. Acetyl CoA is required for fatty acid synthesis in the cyto-
plasm, so there has to be a mechanism for transporting acetyl CoA from the mitochon-
dria to the cytoplasm. This is accomplished using a tricarboxylic acid transporter that
exports citrate. Once in the cytoplasm, citrate has to be reconverted to oxaloacetate and
acetyl CoA and this is accomplished by a cytoplasmic enzyme called ATP- citrate lyase
(Figure 13.15). ATP-citrate lyase doesn’t just catalyze the reverse of the citrate synthase
reaction. The enzyme has to be coupled to hydrolysis of ATP in order to drive the syn-
thesis of “high energy” acetyl CoA in the cytoplasm. The mitochondrial enzyme can
catalyze the same reaction (reversing the citric acid cycle reaction) because the concen-
tration of citrate is so high relative to oxaloacetate (see Figure 13.7). In the cytoplasm,
on the other hand, the steady state concentrations of citrate and oxaloacetate are com-
parable, so coupling to ATP hydrolysis is necessary.

Some species don’t have a mitochondrial version of PEPCK so they have to use an
alternative method of exporting oxaloacetate. The malate-aspartate shuttle is a com-
mon transport system, present even in species that have a mitochondrial PEPCK. A
simplifed version of this shuttle is shown in Figure 13.16. We will describe it in more
detail in Section 14.12.

Oxaloacetate is converted to malate by the reaction catalyzed by malate dehydroge-
nase. This is the same enzyme used in the citric acid cycle. Recall that the equilibrium
concentrations of reactants and products in this reaction result in a very much higher
concentration of malate than oxaloacetate. Thus, a malate transporter is much more
efficient than an oxaloacetate transporter could be.

404 CHAPTER 13 The Citric Acid Cycle

ATP-citrate

lyase

Citrate + ATP + HS-CoA < ± Oxaloacetate + Acetyl CoA + ADP+ Pj

Figure 13.15 ▲

Export of acetyl CoA from mitochondria. Citrate
is exported via the tricarboxylic acid trans-
porter. Citrate is subsequently converted to
acetyl CoA by cytoplasmic ATP-citrate lyase.

Malate is converted back to oxaloacetate by a cytoplasmic version of malate dehy-
drogenase. The net effect is that oxaloacetate from mitochondria can serve as a substrate
for gluconeogenesis as described in the previous chapter.

The other part of the shuttle achieves the same goal by using a mitochondrial
aminotransferase to convert oxaloacetate to aspartate. Aspartate is transported across the
mitochondrial membrane by an aspartate transporter. In the cytoplasm, oxaloacetate can

Figure 13.16 ►

Transport of oxaloacetate via the malate-
aspartate shuttle.

Phosphoenolpyruvate

e o o

""c / NAD© NADH + H

HO — C — H 5

ch 2

Malate

dehydrogenase

(cytoplasmic)

PEPCK

(cytoplasmic)

©o o

i Amino

transferase

c=o < >

o o©

C H;

o o©

©o o

H,N — C — H

r * 1

o /x o®

Malate

Oxaloacetate

Aspartate

Pyruvate

13.5 Reduced Coenzymes Can Fuel Production of ATP

405

be re-formed by the action of a cytoplasmic aminotransferase. As you might guess, this
pathway normally operates in the opposite direction, since the low concentration of ox-
aloacetate in the mitochondria means that the conversion of oxaloacetate to aspartate is
unlikely.

13.5 Reduced Coenzymes Can Fuel Production of ATP

In the net reaction of the citric acid cycle, three molecules of NADH, one molecule of
QH 2 , and one molecule of GTP or ATP are produced for each molecule of acetyl CoA
entering the pathway.

Acetyl CoA + 3 NAD© + Q + GDP (or ADP) + Pj + 2 H 2 0 »

HS-CoA + 3 NADH + QH 2 + GTP (or ATP) + 2 C0 2 + 2 H© (13.15)

As mentioned earlier, NADH and QH 2 can be oxidized by the membrane-asscoci-
ated electron transport chain that is coupled to the the production of ATP. As we will
see when we examine these reactions in Chapter 14, approximately 2.5 molecules of
ATP are generated for each molecule of NADH oxidized to NAD®, and up to 1.5 mol-
ecules of ATP are produced for each molecule of QH 2 oxidized to Q. The complete
oxidation of one molecule of acetyl CoA by the citric acid cycle and subsequent reac-
tions is therefore associated with the production of approximately ten ATP equivalents
(Table 13.2).

The citric acid cycle is the final stage in the catabolism of many major nutrients. It
is the pathway for oxidation of all acetyl CoA molecules produced by the degradation of
carbohydrates, lipids, and amino acids. Having covered glycolysis in Chapter 1 1, we can
now give a complete accounting of the ATP produced from the degradation of one mol-
ecule of glucose.

Recall that glycolysis converts glucose to two molecules of pyruvate with a net gain
of two molecules of ATP. There are two molecules of NADH produced in the reaction
catalyzed by glyceraldehyde 3 -phosphate dehydrogenase. This corresponds to a com-
bined yield of seven ATP equivalents from glycolysis. The conversion of both pyruvate
molecules to acetyl CoA by the pyruvate dehydrogenase complex yields two NADH
molecules, which correspond to about five additional molecules of ATP. When these are
combined with the ATP equivalents from the citric acid cycle via the oxidation of two
molecules of acetyl CoA, the total yield is about 32 molecules of ATP per molecule of
glucose (Figure 13.17).

In bacteria, the two molecules of NADH produced by glycolysis in the cytosol can
be directly reoxidized by the membrane-associated electron transport system in the
plasma membrane. Thus, the theoretical maximum yield from complete oxidation of
glucose (32 ATP equivalents) is achieved in bacteria cells.

In eukaryotic cells, glycolysis produces NADH in the cytosol but the membrane-
associated electron transport complex is located in mitochondria membranes. The reducing
equivalents from cytosolic NADH can be transported into the mitochondrion by shuttle

Table 13.2 Energy production in the citric acid cycle

Reaction

Energy-yielding

product

ATP

equivalents

Isocitrate dehydrogenase

NADH

2.5

a-Ketoglutarate dehydrogenase complex

NADH

2.5

Succinyl-CoA synthetase

GTP or ATP

1.0

Succinate dehydrogenase complex

QH 2

1.5

Malate dehydrogenase

NADH

2.5

Total

10.0

406 CHAPTER 13 The Citric Acid Cycle

Figure 13.17 ►

ATP production from the catabolism of one
molecule of glucose by glycolysis, the citric
acid cycle, and reoxidation of NADH and QH 2 .

The complete oxidation of glucose leads to
the formation of up to 32 molecules of ATP.

ATP

equivalents Glucose

ATP

equivalents

2 ATP «

-> 2 NADH

2 Pyruvate

-> 2 NADH

Substrate level
phosphorylation

2GTP

ATP

2 Acetyl CoA

6 NADH

2 QH 2

Membrane-
associated
electron transport
plus ATP
synthesis
15

4 Total: 32 ATP molecules 28

mechanisms such as the malate-aspartate shuttle described in Section 13.4. The transport
of reducing equivalents of NADH will be described in more detail in Section 14.12.

It’s interesting to compare this pathway (Figure 13.17) for complete oxidation of
glucose to the pentose phosphate cycle described in Section 12.4. That pathway also re-
sults in the complete oxidation of one molecule of glucose. The result is production of
12 NADPH molecules that are equal to 30 ATP equivalents.

13.6 Regulation of the Citric Acid Cycle

Because the citric acid cycle occupies a central position in cellular metabolism, it’s not
surprising to find that the pathway is controlled. Regulation is mediated by allosteric
modulators and by covalent modification of the citric acid cycle enzymes. Flux through
the pathway is further controlled by the supply of acetyl CoA.

As noted earlier, acetyl CoA arises from several sources, including pathways for the
degradation of carbohydrates, lipids, and amino acids. The activity of the pyruvate dehy-
drogenase complex controls the supply of acetyl CoA produced from pyruvate and
hence from the degradation of carbohydrates. In general, substrates of the pyruvate de-
hydrogenase complex activate the complex and products inhibit it. In most species, the
activities of the E 2 and E 3 components of the pyruvate dehydrogenase complex (dihy-
drolipoamide acetyltransferase and dihydrolipoamide dehydrogenase, respectively) are
controlled by simple mass action effects when their products accumulate. The activity of
the acetyltransferase (E 2 ) is inhibited when the concentration of acetyl CoA is high,
whereas the dehydrogenase (E 3 ) is inhibited by a high NADH/NAD© ratio (Figure 13.18).
In general, the inhibitors are likely to be present in high concentrations when energy re-
sources are plentiful, and the activators predominate when energy resources are scarce.

Figure 1 3.1 8 ►

Regulation of the the pyruvate dehydrogenase
complex. Accumulation of the products acetyl
CoA and NADH decreases flux through the
reversible reactions catalyzed by E 2 and E 3 .

Pyruvate + NAD© + HS-CoA

Pyruvate

dehydrogenase

E 1

Dihydrolipoamide

acetyltransferase

Dihydrolipoamide

dehydrogenase

Pyruvate dehydrogenase
complex

NADH + Acetyl CoA + C0 2

13.7 The Citric Acid Cycle Isn’t Always a “Cycle” 407

NAD®, HS-CoA

ADP, Pyruvate NADH, Acetyl CoA

◄ Figure 13.19

Regulation of the mammalian pyruvate dehy-
drogenase complex by phosphorylation of the
component. The regulatory kinase and phos-
phatase are both components of the mam-
malian complex. The kinase is activated by
NADH and acetyl CoA, products of the reac-
tion catalyzed by the pyruvate dehydroge-
nase complex, and inhibited by ADP and the
substrates pyruvate, NAD©, and HS-CoA.

Mammalian (but not prokaryotic) pyruvate dehydrogenase complexes are further
regulated by covalent modification. A protein kinase and a protein phosphatase are as-
sociated with the mammalian multienzyme complex. Pyruvate dehydrogenase kinase
(PDK) catalyzes the phosphorylation of E l5 thereby inactivating the enzyme. Pyruvate
dehydrogenase phosphatase (PDP) catalyzes the dephosphorylation and activation of
pyruvate dehydrogenase (Figure 13.19). Control of E x activity controls the rate of reac-
tion of the entire complex.

Pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase are them-
selves regulated. The kinase is allosterically activated by NADH and acetyl CoA, products of
pyruvate oxidation. The accumulation of NADH and acetyl CoA signals energy availability
and leads to an increase in phosphorylation of the pyruvate dehydrogenase subunit and in-
hibition of the further oxidation of pyruvate. Conversely, pyruvate, NAD®, HS-CoA, and
ADP inhibit the kinase, leading to activation of the pyruvate dehydrogenase subunit.

Three enzymes of the citric acid cycle are regulated: citrate synthase, isocitrate de-
hydrogenase, and the a-ketoglutarate dehydrogenase complex. Citrate synthase cat-
alyzes the first reaction of the citric acid cycle. This would seem to be a suitable control
point for regulation of the entire cycle. ATP inhibits the enzyme in vitro , but significant
changes in ATP concentration are unlikely in vivo ; therefore, ATP may not be a physio-
logical regulator. Some bacterial citrate synthases are activated by a-ketoglutarate and
inhibited by NADH.

Mammalian isocitrate dehydrogenase is allosterically activated by Ca© and ADP
and inhibited by NADH. In mammals, the enzyme is not subject to covalent modifica-
tion. In bacteria, however, isocitrate dehydrogenase is regulated by phosphorylation. We
will discuss this in more detail in Section 13.8.

Although the a-ketoglutarate dehydrogenase complex resembles the pyruvate de-
hydrogenase complex, the enzymes have quite different regulatory features. No kinase or
phosphatase is associated with the a-ketoglutarate dehydrogenase complex. Instead, cal-
cium ions bind to E x of the complex and decrease the K m of the enzyme for a-ketoglutarate,
thereby increasing the rate of formation of succinyl CoA. NADH and succinyl CoA are
inhibitors of the a-ketoglutarate complex in vitro , but it has not been established that
they have a significant regulatory role in living cells.

13.7 The Citric Acid Cycle Isn’t Always a “Cycle”

The citric acid cycle is not exclusively a catabolic pathway for the oxidation of acetyl
CoA. It also plays a central role in metabolism at the intersection of several other path-
ways. Some intermediates of the citric acid cycle are important anabolic precursors in
biosynthesis pathways, and some catabolic pathways produce citric acid cycle interme-
diates. Pathways that are both catabolic and anabolic are said to be amphibolic (Section 10.1).
The citric acid cycle is an excellent example.

408 CHAPTER 13 The Citric Acid Cycle

BOX 13.8 A CHEAP CANCER DRUG?

In the absence of oxygen, the glycolytic pathway terminates
at lactate and the citric acid cycle is not used in the oxidation
of acetyl CoA. Under these conditions, pyruvate dehydroge-
nase is inactivated by phosphorylation. Many cancer cells
grow anaerobically and pyruvate dehydrogenase is not active
in these cells.

The activity of pyruvate dehydrogenase phosphorylase
kinase (PDHK) can be inhibited by dichloroacetate (DCA).
DCA binds to the active site of the enzyme preventing phos-
phorylation of pyruvate dehydrogenase. The net effect of
DCA is activation of pyruvate dehydrogenase and this, in
turn, causes major disruptions in cancer cell metabolism lead-
ing to death of the cancer cells. The chemical has been effec-
tive in a few trial studies with cancer cells in vitro. That’s a
good thing.

Unfortunately, the effectiveness of DCA as a cancer drug
has not been demonstrated in clinical trials. Medical re-
searchers are in a difficult position. The biochemistry is
sound. It makes sense that cancer cells grow anaerobically
(the Warburg effect) and it makes sense that DCA might be
an effective cancer drug based on its ability to inhibit PDHK.
However most physicians are reluctant to prescribe DCA in
the absence of evidence of its effectiveness.

DCA has been around for a long time and it cannot be
patented. This has provoked the claim that major drug com-
panies are conspiring to suppress evidence of DCA’s effec-
tiveness on the grounds that they cannot make any money by
selling DCA. A cottage industry of suppliers has sprung up

on the Internet for people who want to treat themselves with
this cheap “miracle” drug. The Food and Drug Administra-
tion in the United States has been forced to shut down some
websites because they were making unsubstantiated claims
about its ability to cure cancer. There was also concern about
self-medication because high dosages of DCA are toxic.
There’s bound to be more publicity surrounding this compli-
cated issue in the future. The blog Respectful Insolence
(scienceblogs.com/insolence) is a good source of scientific
and medical information on the controversy.

▲ Pyruvate dehydrogenase kinase with dichloroacetate bound at the ac-
tive site. The human ( Homo sapiens ) PDHK is a dimer, only one sub-
unit is shown here. The bound ligands are shown as space-filling
molecules. ADP (top) is bound at the allosteric site, and dichloroac-
etate (left) is bound at the active site. [PDB 2BU8]

As shown in Figure 13.20, citrate, cr-ketoglutarate, succinyl CoA, and oxaloacetate
all lead to biosynthetic pathways. Citrate is part of a pathway for the formation of fatty
acids and steroids. It undergoes cleavage to form acetyl CoA, the precursor of the lipids.
In eukaryotes, this reaction takes place in the cytosol, and citrate must be transported
from the mitochondria to the cytosol to support fatty acid biosynthesis. One major
metabolic fate of a - keto glut ar ate is reversible conversion to glutamate, which can then
be incorporated into proteins or used for the synthesis of other amino acids or nu-
cleotides. We will see in Chapter 17 that a-ketoglutarate pools are important in nitro-
gen metabolism. Succinyl CoA can condense with glycine to initiate the biosynthesis of
porphyrins such as the heme groups of cytochromes. As we saw in the previous chapter,
oxaloacetate is a precursor of carbohydrates formed by gluconeogenesis. Oxaloacetate
also interconverts with aspartate, which can be used in the synthesis of urea, amino
acids, and pyrimidine nucleotides.

When the citric acid cycle functions as a multistep catalyst, only small amounts of
each intermediate are needed to convert large quantities of acetyl CoA to products.
Therefore, the rate at which the citric acid cycle metabolizes acetyl CoA is extremely
sensitive to changes in the concentrations of its intermediates. Thus, citric acid cycle in-
termediates that are removed by entry into biosynthetic pathways must be replenished
by anaplerotic (Greek, “filling up”) reactions. Because the pathway is cyclic, replenishing
any of the cycle intermediates results in a greater concentration of all intermediates. De-
pletion of citric acid cycle intermediates is an example of a cataplerotic reaction. It’s just
as important as the filling up reactions.

The production of oxaloacetate by pyruvate carboxylase is an important anaplerotic
reaction (Figure 13.20). This reaction is also part of the gluconeogenesis pathway
(Section 12.1 A). Pyruvate carboxylase is allosterically activated by acetyl CoA. The ac-
cumulation of acetyl CoA indicates a low concentration of oxaloacetate and a need for

13.8 The Glyoxy late Pathway 409

Carbohydrates

± Alanine

± Fatty acids

Malate

Citrate

steroids

◄ Figure 13.20

Routes leading to and from the citric acid
cycle. Intermediates of the citric acid cycle
are precursors of carbohydrates, lipids, and
amino acids, as well as nucleotides and por-
phyrins. Reactions feeding into the cycle
replenish the pool of cycle intermediates.
Anabolic pathways are colored blue and
catabolic pathways are colored red.

Amino

acids

■> Fumarate

Isocitrate Glutamate

Succinate

u-Ketoglutarate

± Glutamate

Amino acids,
nucleotides

Some amino acids

Propionyl CoA

Porphyrins

Odd-chain fatty acids

more citric acid cycle intermediates. The activation of pyruvate carboxylase supplies ox-
aloacetate for the cycle.

Many species use a variety of different reactions to keep the intake and output of
citric acid cycle intermediates in a delicate balance. For example, many plants and some
bacteria supply oxaloacetate to the citric acid cycle via a reaction catalyzed by phospho-
enolpyruvate carboxylase.

Phosphoenolpyruvate + HCO^ Oxaloacetate + Pj (13.16)

Pathways for degrading some amino acids and fatty acids can contribute succinyl
CoA to the citric acid cycle. The interconversion of oxaloacetate and aspartate and of
a-ketoglutarate and glutamate can either supply or remove intermediates of the cycle.

The interplay of all these reactions — the entry of acetyl CoA from glycolysis and
other sources, the entry of intermediates from catabolic pathways and anaplerotic reac-
tions, and the exit of intermediates to anabolic pathways — means that the citric acid
cycle doesn’t always operate as a simple cycle devoted to oxidizing acetyl CoA. In fact,
most bacteria don’t have all of the classic enzymes of the citric acid cycle so there is no
“cycle” in these species. Instead, the enzymes that are present are used mostly in biosyn-
thesis pathways where the intermediates become precursors for the synthesis of amino
acids and porphyrins (Section 13.9).

13.8 The Glyoxylate Pathway

The glyoxylate pathway is a route that bypasses some of the reactions of the citric acid
cycle. The pathway is named after the two-carbon molecule glyoxylate, an essential

410

CHAPTER 13 The Citric Acid Cycle

intermediate in the pathway. There are only two reactions. In the first reaction, a six-
carbon tricarboxylic acid (isocitrate) is split into a two-carbon molecule (glyoxylate)
and a four-carbon dicarboxylic acid (succinate). This reaction is catalyzed by isocitrate
lyase (Figure 13.21). In the second reaction, the two-carbon glyoxylate molecule com-
bines with a two-carbon acetyl CoA molecule to make a four-carbon dicarboxylic acid
(malate). The enzyme for the second reaction is malate synthase.

The glyoxylate pathway was first discovered in bacteria. Subsequently it was found
in plants and later in fungi, protists, and some animals. The pathway is often called the
glyoxylate shunt, the glyoxylate bypass, or the glyoxylate cycle. The glyoxylate pathway
provides an anabolic alternative for the metabolism of acetyl CoA, leading to the formation
of glucose from acetyl CoA via four-carbon compounds. Cells that contain glyoxylate
pathway enzymes can synthesize all their required carbohydrates from any substrate
that is a precursor of acetyl CoA. For example, yeast can grow on ethanol because yeast
cells can oxidize ethanol to form acetyl CoA, which can be metabolized via the glyoxylate
pathway to form malate. Similarly, many bacteria use the glyoxylate pathway to sustain
growth on acetate, which can be incorporated into acetyl CoA in a reaction catalyzed by
acetyl CoA synthetase.

AMP # PPj

ATP t O

H,C — COO 0 + HS-CoA — ^ ^ — » H,C — C— S-CoA (13.17)

Acetate synthetase Acetyl CoA

The glyoxylate pathway is a fundamental metabolic pathway in bacteria, protists,
fungi, and plants. It is especially active in oily seed plants. In these plants, stored seed
oils (triacylglycerols) are converted to carbohydrates that provide fuel during germina-
tion. In contrast, genes for the two enzymes of the pathway are present in most animals
but the pathway is not actively used. Consequently, in humans acetyl CoA does not
serve as the precursor for the net formation of either pyruvate or oxaloacetate; there-
fore, acetyl CoA is not a carbon source for the net production of glucose. (The carbon
atoms of acetyl CoA are incorporated into oxaloacetate by the reactions of the citric
acid cycle, but for every two carbon atoms incorporated, two other carbon atoms are
released as C0 2 .)

The glyoxylate pathway can be regarded as a shunt within the citric acid cycle, as
shown in Figure 13.21. The two reactions provide a bypass around the C0 2 -producing
reactions of the citric acid cycle. No carbon atoms of the acetyl group of acetyl CoA are
released as C0 2 during operation of the glyoxylate shunt, and the net formation of a
four-carbon molecule from two molecules of acetyl CoA supplies a precursor that can
be converted to glucose by gluconeogenesis. Succinate is oxidized to malate and ox-
aloacetate by the citric acid cycle to maintain the catalytic amounts of citric acid cycle
intermediates. You can think of the glyoxylate shunt as part of a cycle that includes the
upper portion of the citric acid cycle. In this case, the net reaction includes the forma-
tion of oxaloacetate for gluconeogenesis and the cyclic oxidation of succinate. Two mol-
ecules of acetyl CoA are consumed.

2 Acetyl CoA + 2 NAD© + Q + 3 H 2 0 »

Oxaloacetate + 2 HS-CoA + 2 NADH + QH 2 + 4 H© (13.18)

In eukaryotes, the operation of the glyoxylate cycle requires the transfer of metabo-
lites between the mitochondria, where the citric acid cycle enzymes are located, and the
cytosol, where isocitrate lyase and malate synthase are found. Thus, the actual pathway
is more complicated than the diagram in Figure 13.21. In plants, the glyoxylate pathway
enzymes are localized to a special membrane-bound organelle called the glyoxysome.
Glyoxysomes contain some special versions of the citric acid cycle enzymes, but some
metabolites still have to be transferred between compartments in order for the pathway
to operate as a cycle.

13.8 The Glyoxylate Pathway 41 1

j©

HO — C — H

C H 2

COO 0

L-Malate

Fumarase

H.O'

CH 3

c=o
I

S-CoA
Acetyl CoA

Malate synthase

,©

H.O

COO (

n HS-CoA,H

oh 2

HO— C — COO 1

ch 2

coo°

Citrate

Aconitase

coo 0

◄ Figure 13.21

Glyoxylate pathway. Isocitrate lyase and
malate synthase are the two enzymes of the
pathway. When the pathway is functioning,
the acetyl carbon atoms of acetyl CoA are
converted to malate rather than oxidized to
CO 2 . Malate can be converted to oxaloac-
etate, which is a precursor in gluconeogene-
sis. The succinate produced in the cleavage
of isocitrate is oxidized to oxaloacetate to re-
place the four-carbon compound consumed
in glucose synthesis.

H — C

C — H

1 ©

coo°

Fumarate

Succinate

CH,

C> H

coo'

H — C — COO'

HO— C — H

,©

COO

Succinyl CoA

In bacteria, the glyoxylate pathway is often used to replenish citric acid cycle
metabolites that are diverted into a number of biosynthesis pathways. Since all of the re-
actions take place in the cytosol in bacteria, it is important to regulate the flow of
metabolites. The key regulated enzyme is isocitrate dehydrogenase. Its activity is regu-
lated by covalent modification. Kinase-catalyzed phosphorylation of a serine residue
abolishes isocitrate dehydrogenase activity. In the dephosphorylated form of the en-
zyme, the serine residue forms a hydrogen bond with a carboxylate group of isocitrate.
Phosphorylation inhibits enzyme activity by causing electrostatic repulsion of the sub-
strate rather than by causing an R-to-T conformational change (Figure 13.22). The
same protein molecule that contains the kinase activity also has a separate domain with
phosphatase activity that catalyzes hydrolysis of the phosphoserine residue, reactivating
isocitrate dehydrogenase.

The kinase and phosphatase activities are reciprocally regulated; isocitrate,
oxaloacetate, pyruvate, and the glycolytic intermediates 3-phosphoglycerate and phos-
phoenolpyruvate allosterically activate the phosphatase and inhibit the kinase

▲ Figure 13.22

Phosphorylated and dephosphorylated forms of
E. coli iso citrate dehydrogenase, (a) The de-
phosphorylated enzyme is active; isocitrate
binds to the active site. [PDB5ICD] (b) The
phosphorylated enzyme is inactive because
the negatively charged phosphoryl group
(red) electrostatically repels the substrate,
preventing it from binding. [PDB4ICD]

412 CHAPTER 13 The Citric Acid Cycle

Figure 13.23 ►

Regulation of E. coli isocitrate dehydrogenase
by covalent modification. A bifunctional en-
zyme catalyzes phosphorylation and dephos-
phorylation of isocitrate dehydrogenase. The
two activities of the bifunctional enzyme are
reciprocally regulated allosterically by inter-
mediates of glycolysis and the citric acid
cycle.

Isocitrate

ATP

- 1

ADP

Isocitrate j Bifunctional I Isocitrate
dehydrogenase kinase/phosphatase I dehydrogenase I

( Isocitrate \_ft)
(dehydrogenase

active

u-Ketoglutarate

inactive

FUO

Isocitrate

Oxaloacetate

Pyruvate

3-Phosphoglycerate

Phosphoenolpyruvate

(Figure 13.23). Thus, when the concentrations of glycolytic and citric acid cycle inter-
mediates in E. coli are high, isocitrate dehydrogenase is active. When phosphorylation
abolishes the activity of isocitrate dehydrogenase, isocitrate is diverted to the glyoxylate
pathway.

13.9 Evolution of the Citric Acid Cycle

The reactions of the citric acid cycle were first discovered in mammals and many of the
key enzymes were purified from liver extracts. As we have seen, the citric acid cycle can
be viewed as the end stage of glycolysis because it results in the oxidation of acetyl CoA
produced as one of the products of glycolysis. However, there are many organisms that
do not encounter glucose as a major carbon source and the production of ATP equiva-
lents via glycolysis and the citric acid cycle is not an important source of metabolic
energy in such species.

We need to examine the function of the citric acid cycle enzymes in bacteria in
order to understand their role in simple single-celled organisms. These roles might
allow us to deduce the pathways that could have existed in the primitive cells that even-
tually gave rise to complex eukaryotes. Fortunately, the sequences of several hundred
prokaryotic genomes are now available as a result of the huge technological advances in
recombinant DNA technology and DNA sequencing methods. We can now examine the
complete complement of metabolic enzymes in many diverse species of bacteria and
ask whether they possess the pathways that we have discussed in this chapter. These
analyses are greatly aided by developments in the fields of comparative genomics,
molecular evolution, and bioinformatics.

Most species of bacteria do not have a complete citric acid cycle. The most com-
mon versions of an incomplete cycle include part of the left-hand side. This short linear
pathway leads to production of succinate or succinyl CoA or a-ketoglutarate by a re-
ductive process using oxaloacetate as a starting point. This reductive pathway is the
reverse of the traditional cycle that functions in the mitochondria of eukaryotes. In ad-
dition, many species of bacteria also have enzymes from part of the right-hand side of
the citric acid cycle, especially citrate synthase and aconitase. This allows them to syn-
thesize citrate and isocitrate from oxaloacetate and acetyl CoA. The presence of a forked
pathway (Figure 13.24) results in the synthesis of all the precursors of amino acids,
porphyrins, and fatty acids.

There are hundreds of diverse species of bacteria that can survive and grow in the
complete absence of oxygen. Some of these species are obligate anaerobes — for them,
oxygen is a lethal poison! Others are facultative anaerobes — they can survive in oxygen
free environments as well as oxygen- rich environments. E. coli is one example of a species
that can survive in both types of environment. When growing anaerobically, E. coli uses a
forked version of the pathway to produce the necessary metabolic precursors and avoid
the accumulation of reducing equivalents that cannot be reoxidized by the oxygen
requiring electron transport system. Bacteria such as E. coli can grow in environments
where acetate is the only source of organic carbon. In this case, they employ the glyoxy-
late pathway to convert acetate to malate and oxaloacetate for glucose synthesis.

13.9 Evolution of the Citric Acid Cycle

413

Oxaloacetate

Malate dehydrogenase

Acetyl CoA

Citrate synthase

■>

Citrate

Aconitase

Oxidative

pathway

Reductive

pathway

Malate

Fumarase

Fumarate

Fumarate reductase

Malate

synthase

Isocitrate

Acetyl CoA

Glyoxylate

Isocitrate

dehydrogenase

Succinate

dehydrogenase

k C0 2
a-Ketoglutarate

Succinate

Isocitrate

lyase

Succinyl CoA synthetase
Succinyl CoA: acetoacetate
CoA transferase

Succinyl CoA

a-Ketoglutarate QQ ?
Dehydrogenase ^
a-Ketoglutarate:

Ferredoxin oxidoreductase N ,

a-Ketoglutarate

◄ Figure 13.24

Forked pathway found in many species of bac-
teria. The left-hand side of the fork is a re-
ductive pathway leading to the synthesis of
succinate or a-ketoglutarate in reactions
that proceed in the reverse direction from
those in the classic citric acid cycle. The
right-hand branch is an oxidative pathway
similar to the first few reactions of the clas-
sic citric acid cycle.

The first living cells arose in an oxygen-free environment over three billion years
ago. These primitive cells undoubtedly possessed most of the enzymes that intercon-
verted acetate, pyruvate, citrate, and oxaloacetate, since these enzymes are present in
most modern bacteria. The development of the main branches of the forked pathway
possibly began with the evolution of malate dehydrogenase from a duplication of the
lactate dehydrogenase gene. Aconitase and isocitrate dehydrogenase evolved from en-
zymes that are used in the synthesis of leucine (isopropylmalate dehydratase and iso-
propylmalate dehydrogenase, respectively). (Note that the leucine biosynthesis pathway
is more ubiquitous and more primitive than the citric acid cycle.)

Extension of the reductive branch continued with the evolution of fumarase from
aspartase. Aspartase is a common bacterial enzyme that synthesizes fumarate from
L-aspartate. L-aspartate, in turn, is synthesized by amination of oxaloacetate in a reac-
tion catalyzed by aspartate transaminase (Section 17.3). It is likely that primitive cells
used the pathway oxaloacetate — > aspartate — > fumarate to produce fumarate before the
evolution of malate dehydrogenase and fumarase. The reduction of fumarate to succi-
nate is catalyzed by fumarate reductase in many bacteria. The evolutionary origin of
this complex enzyme is highly speculative but at least one of the subunits is related to
another enzyme of amino acid metabolism. Succinate dehydrogenase, the enzyme that
preferentially catalyzes the reverse reaction in the citric acid cycle, is likely to have
evolved later on from fumarate reductase via a gene duplication event.

The synthesis of a-ketoglutarate can occur in either branch of the forked pathway.
The reductive branch uses a-ketoglutarate:ferredoxin oxidoreductase, an enzyme found
in many species of bacteria that don’t have a complete citric acid cycle. The reaction cat-
alyzed by this enzyme is not readily reversible. With the evolution of a-ketoglutarate de-
hydrogenase the two forks can be joined to create a cyclic pathway. It is clear that
a-ketoglutarate dehydrogenase and pyruvate dehydrogenase share a common ancestor
and it is likely that this was the last enzyme to evolve.

Some bacteria have a complete citric acid cycle but it is used in the reductive direction
to fix C0 2 in order to build more complex organic molecules. This could have been one of
the selective pressures leading to a complete pathway. The cycle requires a terminal elec-
tron acceptor to oxidize NADH and QH 2 when it operates in the more normal oxidative
direction seen in eukaryotes. Originally, this terminal electron acceptor was sulfur or vari-
ous sulfates, and these reactions still occur in many anaerobic bacterial species. Oxygen
levels began to rise about 2.5 billion years ago with the evolution of photosynthesis reac-
tions in cyanobacteria. Some bacteria, notably proteobacteria, exploited the availability of

414

CHAPTER 13 The Citric Acid Cycle

oxygen when the membrane-associated electron transport reactions evolved. One species
of proteobacteria entered into a symbiotic relationship with a primitive eukaryotic cell
about two billion years ago. This led to the evolution of mitochondria and the modern
versions of the citric acid cycle and electron transport in eukaryotes.

The evolution of the citric acid cycle pathway involved several of the pathway evo-
lution mechanisms discussed in Chapter 10. There is evidence for gene duplication,
pathway extension, retro -evolution, pathway reversal, and enzyme theft.

Summary

1. The pyruvate dehydrogenase complex catalyzes the oxidation of
pyruvate to form acetyl CoA and C0 2 .

2. For each molecule of acetyl CoA oxidized via the citric acid cycle,
two molecules of C0 2 are produced, three molecules of NAD®
are reduced to NADH, one molecule of Q is reduced to QH 2 and
one molecule of GTP is generated from GDP + Pj (or ATP from
ADP + P^ depending on the species).

3. The eight enzyme- catalyzed reactions of the citric acid cycle can
function as a multistep catalyst.

4. In eukaryotic cells, pyruvate must be imported into the mitochon-
dria by a specific transporter before it can serve as a substrate for
the pyruvate dehydrogenase reaction.

5. Oxidation of the reduced coenzymes generated by the citric acid cycle
leads to the formation of about 10 ATP molecules per molecule of

acetyl CoA entering the pathway, for a total of about 32 ATP mol-
ecules per complete oxidation of 1 molecule of glucose.

6. The oxidation of pyruvate is regulated at the steps catalyzed by
the pyruvate dehydrogenase complex, isocitrate dehydrogenase,
and the a-ketoglutarate dehydrogenase complex.

7. In addition to its role in oxidative catabolism, the citric acid cycle
provides precursors for biosynthetic pathways. Anaplerotic reac-
tions replenish cycle intermediates.

8. The glyoxylate cycle, a modification of the citric acid cycle, allows
many organisms to use acetyl CoA to generate four- carbon inter-
mediates for gluconeogenesis.

9. The citric acid cycle probably evolved from the more
primitive forked pathway found in many modern species of
bacteria.

Problems

1. (a) The citric acid cycle converts one molecule of citrate to one

molecule of oxaloacetate, which is required for the cycle to
continue. If other cycle intermediates are depleted by being
used as precursors for amino acid biosynthesis, can a net syn-
thesis of oxaloacetate occur from acetyl CoA via the enzymes
of the citric acid cycle?

(b) How can the cycle continue to function if insufficient oxalo-
acetate is present?

2. Fluoroacetate, a very toxic molecule that blocks the citric acid
cycle, has been used as a rodent poison. It is converted enzymati-
cally in vivo to fluoroacetyl CoA, which is then converted by the
action of citrate synthase to 2R,3S-fluorocitrate, a potent competi-
tive inhibitor of the next enzyme in the pathway. Predict the effect
of fluoroacetate on the concentrations of the intermediates in the
citric acid cycle. How can this blockage of the cycle be overcome?

3. Calculate the number of ATP molecules generated by the follow-
ing net reactions of the citric acid cycle. Assume that all NADH
and QH 2 are oxidized to yield ATP, pyruvate is converted to acetyl
CoA, and the malate-aspartate shuttle is operating.

(a) 1 Pyruvate » 3 C0 2

(b) Citrate > Oxaloacetate + 2 C0 2

4. When one molecule of glucose is completely oxidized to six mole-
cules of C0 2 under the conditions in Problem 3, what percentage
of ATP is produced by substrate level phosphorylation?

5. The disease beriberi, which results from a dietary deficiency of vi-
tamin Bi (thiamine), is characterized by neurologic and cardiac

symptoms, as well as increased levels of pyruvate and a-ketoglu-
tarate in the blood. How does a deficiency of thiamine account for
the increased levels of pyruvate and a-ketoglutarate

6. In three separate experiments, pyruvate labeled with 14 C at C- 1, at
C-2, or at C-3 is metabolized via the pyruvate dehydrogenase
complex and the citric acid cycle. Which labeled pyruvate mole-
cule is the first to yield 14 C0 2 ? Which is the last to yield 14 C0 2 ,
and how many turns of the cycle are required to release all of the
labeled carbon atoms as 14 C0 2 ?

7. Patients in shock experience decreased delivery of 0 2 to tissues, de-
creased activity of the pyruvate dehydrogenase complex, and in-
creased anaerobic metabolism. Excess pyruvate is converted to lactate,
which accumulates in tissues and in the blood, causing lactic acidosis.

(a) Since 0 2 is not a reactant or product of the citric acid cycle,
why do low levels of 0 2 decrease the activity of the pyruvate
dehydrogenase complex?

(b) To alleviate lactic acidosis, shock patients are sometimes
given dichloroacetate, which inhibits pyruvate dehydroge-
nase kinase. How does this treatment affect the activity of the
pyruvate dehydrogenase complex?

8. A deficiency of a citric acid cycle enzyme in both mitochondria
and the cytosol of some tissues (e.g., blood lymphocytes) results
in severe neurological abnormalities in newborns. The disease is
characterized by excretion in the urine of abnormally large
amounts of cr-ketoglutarate, succinate, and fumarate. What en-
zyme deficiency would lead to these symptoms?

Problems 415

9. Acetyl CoA inhibits dihydrolipoamide acetyltransferase (E 2 of the
pyruvate dehydrogenase complex) but activates the pyruvate de-
hydrogenase kinase component of the pyruvate dehydrogenase
complex. How are these two different actions of acetyl CoA con-
sistent with the overall regulation of the complex?

10 . Pyruvate dehydrogenase complex deficiency is a disease that
results in various metabolic and neurological effects. Pyruvate
dehydrogenase complex deficiency can cause lactic acidosis in
affected children. Other clinical symptoms include increased con-
centrations of pyruvate and alanine in the blood. Explain the
increase in the levels of pyruvate, lactate, and alanine in individu-
als with pyruvate dehydrogenase complex deficiency.

11. In response to a signal for contraction and the resulting increased
need for ATP in vertebrate muscle, Ca© is released into the cy-
tosol from storage sites in the endoplasmic reticulum. How does
the citric acid cycle respond to the influx of Ca© in satisfying the
increased need for cellular ATP?

12. (a) The degradation of alanine yields pyruvate, and the degrada-

tion of leucine yields acetyl CoA. Can the degradation of
these amino acids replenish the pool of citric acid cycle inter-
mediates?

(b) Fats (triacylglycerols) stored in adipose tissue are a signifi-
cant source of energy in animals. Fatty acids are degraded to
acetyl CoA, which activates pyruvate carboxylase. How does the
activation of this enzyme help recover energy from fatty acids?

13 . Amino acids resulting from the degradation of proteins can be
further metabolized by conversion to intermediates of the citric
acid cycle. If the degradation of a labeled protein leads to the

following labeled amino acids, write the structure of the first
intermediate of the citric acid cycle into which these amino
acids would be converted and identify the labeled carbon in
each case.

(a) COO 0

ch 2

14 ch 2

(b) CH 3 (c)

HjN 1 — 14 CH — COO 0 14 COO°

Alanine J.

CH 2

H 3 N — CH — COO 0

H 3 N— CH — COO 0

Glutamate

Aspartate

14 . (a) How many molecules of ATP are eventually generated when

two molecules of acetyl CoA are converted to four molecules
of C0 2 via the citric acid cycle? (Assume NADH 2.5 ATP and
QH 2 -1.5ATP) How many molecules of ATP are generated
when two molecules of acetyl CoA are converted to oxaloac-
etate in the glyoxylate cycle?

(b) How do the yields of ATP relate to the primary functions of
the two pathways?

15 . The activities of PFK-2 and fructose 2,6-frisphosphatase are
contained in a bifunctional protein that effects tight control over
glycolysis and gluconeogenesis through the action of fructose
2,6-hisphosphate. Describe another protein that contains kinase
and phosphatase activities in a single protein molecule. What
pathways does it control?

41 6 CHAPTER 13 The Citric Acid Cycle

Selected Readings

Pyruvate Dehydrogenase Complex

Harris, R. A., Bowker-Kinley, M. M., Huang, B.,
and Wu, R (2002). Regulation of the activity of the
pyruvate dehydrogenase complex. Advances in
Enzyme Regulation 42:249-259.

Knoechel, T. R., Tucker, A. D., Robinson, C. M.,
Phillips, C., Taylor, W., Bungay, P. J., Kasten, S. A.,
Roche, T. E., and Brown, D. G. (2006). Regulatory
roles of the N-terminal domain based on crystal
structures of human pyruvate dehydrogenase kinase 2
containing physiological and synthetic ligands.
Biochem. 45:402-415.

Maeng, C.-Y., Yazdi, M. A., Niu, X.-D., Lee, H. Y., and
Reed, L. J. (1994). Expression, purification, and char-
acterization of the dihydrolipoamide dehydroge-
nase-binding protein of the pyruvate dehydroge-
nase complex from Saccharomyces cerevisiae.
Biochem. 33:13801-13807.

Mattevi, A., Obmolova, G., Schulze, E., Kalk, K. H.,
Westphal, A. H., de Kok, A., and Hoi, W. G. J.
(1992). Atomic structure of the cubic core of the
pyruvate dehydrogenase multienzyme complex.
Science 255:1544-1550.

Reed, L. J., and Hackert, M. L. (1990). Structure-
function relationships in dihydrolipoamide acyl-
transferases. /. Biol. Chem. 265:8971-8974.

Citric Acid Cycle

Beinert, H., and Kennedy, M. C. (1989). Engineer-
ing of protein bound iron-sulfur clusters. Eur. J.
Biochem. 186:5-15.

Gruer, M. J., Artymiuk, P. J., and Guest, J. R. (1997).
The aconitase family: three structural variations
on a comon theme. Trends Biochem. Sci. 22:3-6.

Hurley, J. H., Dean, A. M., Sohl, J. L., Koshland,

D. E., Jr., and Stroud, R. M. (1990). Regulation of
an enzyme by phosphorylation at the active site.
Science 249:1012-1016.

Kay, J., and Weitzman, P. D. J., eds. (1987). Krebs’
Citric Acid Cycle — Haifa Century and Still Turning
(London: The Biochemical Society).

Krebs, H. A., and Johnson, W. A. (1937). The role
of citric acid in intermediate metabolism in ani-
mal tissues. Enzymologia 4:148-156.

McCormack, J. G., and Denton, R. M. (1988). The
regulation of mitochondrial function in mam-
malian cells by Ca® ions. Biochem. Soc. Trans.
109:523-52 7.

Remington, S. J. (1992). Mechanisms of citrate
synthase and related enzymes (triose phosphate
isomerase and mandelate racemase). Curr.

Opin. Struct. Biol. 2:730-735.

Williamson, J. R., and Cooper, R. H. (1980). Regu-
lation of the citric acid cycle in mammalian sys-
tems. FEBS Lett. 117 (Suppl.):K73-K85.

Wolodko, W. T., Fraser, M. E., James, M. N. G., and
Bridger, W. A. (1994). The crystal structure of suc-
cinyl-CoA synthetase from Escherichia coli at 2.5-A
resolution./. Biol. Chem. 269:10883-10890.

Yankovskaya, V., Horsefield, R., Tornroth, S.,
Luna-Chavez, C., Miyoshi, H., Leger, C., Byrne, B.,
Cecchini, G. and Iwata, S. (2003). Architecture of
succinate dehydrogenase and reactive oxygen
species generation. Science 299:700-704.

Glyoxylate Cycle

Beevers, H. (1980). The role of the glyoxylate
cycle. In The Biochemistry of Plants: A Compre-
hensive Treatise , Vol. 4, P. K. Stumpf and E. E.

Conn, eds. (New York: Academic Press),
pp. 117-130.

o c

_ o

° o o o

° o

o o

° c

o o

Electron Transport
and ATP Synthesis

W e now come to one of the most complicated metabolic pathways encoun-
tered in biochemistry — the membrane-associated electron transport
system coupled to ATP synthesis. The role of this pathway is to convert re-
ducing equivalents into ATP. We usually think of reducing equivalents as products
of glycolysis and the citric acid cycle since the oxidation of glucose and acetyl CoA is
coupled to the reduction of NAD® and Q. In this chapter we learn that the subsequent
reoxidation of NADH and QH 2 results in the passage of electrons through a membrane-
associated electron transport system where the energy released can be saved through the
phosphorylation of ADP to ATP. The electrons are eventually passed to a terminal elec-
tron acceptor. This terminal electron acceptor is usually molecular oxygen (0 2 ) and this
is why the overall process is often called oxidative phosphorylation.

The combined pathway of electron transport and ATP synthesis involves numer-
ous enzymes and coenzymes. It also depends absolutely on the presence of a mem-
brane compartment since one of the key steps in coupling electron transport to ATP
synthesis involves the creation of a pH gradient across a membrane. In eukaryotes the
membrane is the inner mitochondrial membrane and in prokaryotes it is the plasma
membrane.

We begin this chapter with an overview of the thermodynamics of a proton gradi-
ent and how it can drive ATP synthesis. We then describe the structure and function of
the membrane-associated electron transport complexes and the ATP synthase complex.
We conclude with a description of other terminal electron acceptors and a brief discus-
sion of some enzymes involved in oxygen metabolism. Chapter 15 describes the similar
membrane- associated electron transport and ATP synthesis pathway that operates during
photosynthesis.

According to the chemiosmotic hy-
pothesis of oxidative and photosyn-
thetic phosphorylation proposed by
Mitchell the linkage between electron
transport and phosphorylation occurs
not because of hypothetical energy-
rich chemical intermediates as in the
orthodox view, but because oxido-
reduction and adenosine triphosphate
(ATP) hydrolysis are each separately
associated with the net translocation
of a certain number of electrons in
one direction and the net transloca-
tion of the same number of hydrogen
atoms in the opposite direction across
a relatively ion-, acid-, and base-
impermeable coupling membrane.

P Mitchell, and J. Moyle, (1965)

Top: Sunflowers, cheetahs, and mushrooms all use the same mechanism to make ATP using a proton gradient.

417

418

CHAPTER 14 Electron Transport and ATP Synthesis

The coenzymes mentioned in this chap-
ter are described in detail in Chapter 7:
NAD + , Section 7.4; ubiquinone,

Section 7.15; FMN and FAD, Section 7.5;
iron-sulfur clusters, Section 7.1; and
cytochromes, Section 7.17.

14.1 Overview of Membrane-associated
Electron Transport and ATP Synthesis

Membrane-associated electron transport requires several enzyme complexes embedded
in a membrane. We will start by examining the pathway that occurs in mitochondria
and later we will look at the common features of the prokaryotic and eukaryotic sys-
tems. The two processes of membrane- associated electron transport and ATP synthesis
are coupled — neither process can occur without the other.

In the common pathway, electrons are passed from NADH to the terminal electron
acceptor. There are many different terminal electron acceptors but we are mostly inter-
ested in the pathway found in eukaryotic mitochondria where molecular oxygen (0 2 ) is
reduced to form water. As electrons pass along the electron transport chain from NADH
to 0 2 the energy they release is used to transfer protons from inside the mitochondrion
to the intermembrane space between the double membranes. This proton gradient is
used to drive ATP synthesis in a reaction catalyzed by ATP synthase (Figure 14.1). A very
similar system operates in bacteria.

As mentioned above, the entire mitochondrial pathway is often called oxidative
phosphorylation because, historically, the biochemical puzzle was to explain the linkage
between oxygen uptake and ATP synthesis. You will also see frequent references to “res-
piration” and “respiratory electron transport.” These terms also refer to the pathway
that exploits oxygen as the terminal electron acceptor.

v Figure 14.1

Overview of membrane-associated electron
transport and ATP synthesis in mitochondria.

A proton concentration gradient is produced
from reactions catalyzed by the electron
transport chain. Protons are translocated
across the inner mitochondrial membrane
from the matrix to the intermembrane space
as electrons from reduced substrates flow
through the complexes. The free energy
stored in the proton concentration gradient
is utilized when protons flow back across
the membrane via ATP synthase; their reen-
try is coupled to the conversion of ADP and
Pi to ATP.

14.2 The Mitochondrion

Much of the aerobic oxidation of biomolecules in eukaryotes takes place in the mito-
chondrion. This organelle is the site of the citric acid cycle and fatty acid oxidation,
both of which generate reduced coenzymes. The reduced coenzymes are oxidized by the
electron transport complexes embedded in the mitochondrial membranes. The struc-
ture of a typical mitochondrion is shown in Figure 14.2.

The number of mitochondria in cells varies dramatically. Some unicellular algae
contain only one mitochondrion whereas the cell of the protozoan Chaos chaos con-
tains half a million mitochondria. A mammalian liver cell contains up to 5000 mitochondria.
The number of mitochondria is related to the overall energy requirements of the cell.
White muscle tissue, for example, relies on anaerobic glycolysis for its energy needs and
it contains relatively few mitochondria. The rapidly contracting but swiftly exhausted
jaw muscles of the alligator are an extreme example of white muscle. Alligators can snap
their jaws with astonishing speed and force but cannot continue this motion beyond a

14.2 The Mitochondrion 419

very few repetitions. By contrast, red muscle tissue has many mitochondria. The cells of
the flight muscles of migratory birds are an example of red muscle cells. These muscles
must sustain substantial and steady outputs of power and this power requires prodi-
gious amounts of ATP.

Mitochondria vary greatly in size and shape among different species, in different
tissues, and even within a cell. A typical mammalian mitochondrion has a diameter of
0.2 to 0.8 //m and a length of 0.5 to 1.5 //m — this is about the size and shape of an E. coli
cell. (Recall from Chapter 1 that mitochondria are descendants of bacteria cells that en-
tered into a symbiotic relationship with a primitive eukaryotic cell.)

Mitochondria are separated from the cytoplasm by a double membrane. The two
membranes have markedly different properties. The outer mitochondrial membrane
has few proteins. One of these proteins is the transmembrane protein porin (Section
9. 11 A) that forms channels allowing free diffusion of ions and water-soluble metabo-
lites with molecular weights less than 10,000. In contrast, the inner mitochondrial
membrane is very rich in protein with a protein-to-lipid ratio of about 4:1 by mass.
This membrane is permeable to uncharged molecules such as water, 0 2 , and C0 2 but it
is a barrier to protons and larger polar and ionic substances. These polar substances
must be actively transported across the inner membrane using specific transport pro-
teins such as pyruvate translocase (Section 13.4). The entry of anionic metabolites into
the negatively charged interior of a mitochondrion is energetically unfavorable. Such
metabolites are usually exchanged for other anions from the interior or are accompa-
nied by protons flowing down the concentration gradient that is generated by the elec-
tron transport chain.

The inner membrane is often highly folded resulting in a greatly increased surface
area. The folds are called cristae. The expansion and folding of the inner membrane also
creates a greatly expanded intermembrane space (Figure 14.2a). Since the outer membrane
is freely permeable to small molecules, the intermembrane space has about the same
composition of ions and metabolites as the cytosol that surrounds the mitochondrion.

The contents of the matrix include the pyruvate dehydrogenase complex, the en-
zymes of the citric acid cycle (except for the succinate dehydrogenase complex, which is
embedded in the inner membrane), and most of the enzymes that catalyze fatty acid ox-
idation. The protein concentration in the matrix is very high (approaching 500 mg ml -1 ).
Nevertheless, diffusion is only slightly less rapid than in the cytosol (Section 2.3b).

◄ Figure 14.2

Structure of the mitochondrion. The outer
mitochondrial membrane is freely permeable
to small molecules but the inner membrane
is impermeable to polar and ionic substances.
The inner membrane is highly folded and
convoluted, forming structures called cristae.
The protein complexes that catalyze the
reactions of membrane-associated electron
transport and ATP synthesis are located in
the inner membrane, (a) Illustration.

(b) Electron micrograph: longitudinal section
from bat pancreas cell.

▲ Alligator jaw muscles. You’re probably safe
after this alligator has already snapped at
you several times and missed. (If you trust
your biochemistry textbook.)

V, v *

▲ Canada geese. If you had more mitochon-
dria in your muscle cells you might be able
to fly to a warmer climate for the winter.

420 CHAPTER 14 Electron Transport and ATP Synthesis

BOX 14.1 AN EXCEPTION TO EVERY RULE

One of the most fascinating things about biology is that there are very few universal
rules. We can propose certain general principles that apply in most cases but there
are almost always a few examples that don’t fit. For example, we can say that eukary-
otic cells contain mitochondria as a general rule but we know of some species that
don’t have mitochondria.

One of the “rules” that seemed valid was that all animal cells had mitochondria
and they all require oxygen. Now there’s even an exception to that rule. Some small
microscopic animals of the phylum Loricifera live in deep ocean basins where there
is no light and the nearly salt-saturated water is devoid of oxygen. They are inca-
pable of aerobic oxidation and their cells have no mitochondria.

► Spinoloricus sp., an anaerobic animal.

▲ Peter Mitchell (1920-1992). Mitchell was
awarded the Nobel Prize in Chemistry in
1978 “for his contribution to the understand
ing of biological energy transfer through the
formulation of the chemiosmotic theory.”

In 1963 Mitchell resigned from his position
at Edinburgh University in Scotland and in
1965 he set up a private research institute
with his long-time friend and collaborator,
Jennifer Moyle. They continued to work on
bioenergetics in a laboratory in Mitchell’s
home, Glynn House, in Cornwall (UK).

KEY CONCEPT

Chemiosmotic theory states that the
energy from the oxidation-reduction
reactions of electron transport is used to
create a proton gradient across the
membrane and the resulting protonmotive
force is used in the synthesis of ATP.

The matrix also contains metabolites and inorganic ions and a pool of NAD® and
NADP® that remains separate from the pyridine nucleotide coenzymes of the cytosol.
Mitochondrial DNA and all of the enzymes required for DNA replication, transcrip-
tion, and translation are located in the matrix. Mitochondrial DNA contains many of
the genes that encode the electron transport proteins (see Figure 14.19 ).

14.3 The Chemiosmotic Theory and Protonmotive Force

Before considering the individual reactions of oxidative phosphorylation we will exam-
ine the nature of the energy stored in a proton concentration gradient. The
chemiosmotic theory is the concept that a proton concentration gradient serves as the en-
ergy reservoir that drives ATP formation. The essential elements of this theory were
originally formulated by Peter Mitchell in the early 1960s. At the time, the mechanism
by which cells carry out oxidative phosphorylation was the subject of intensive research
and much controversy. The pathway linking oxidation reactions to the phosphorylation
of ADP was not known and many early attempts to identify a “high energy” phosphory-
lated metabolite that could transfer a phosphoryl group to ADP had ended in failure.
Today, thanks to decades of work by many scientists, the formation and dissipation of
ion gradients are acknowledged as a central motif in bioenergetics. Mitchell was
awarded the Nobel Prize in Chemistry in 1978 for his contribution to our understand-
ing of bioenergetics.

A. Historical Background: The Chemiosmotic Theory

By the time Mitchell proposed the chemiosmotic theory, much information had accu-
mulated on the oxidation of substrates and the cyclic oxidation and reduction of mito-
chondrial electron carriers. In 1956 Britton Chance and Ronald Williams had shown
that when intact isolated mitochondria are suspended in phosphate buffer they oxidize
substrates and consume oxygen only when ADP is added to the suspension. In other
words, the oxidation of a substrate must be coupled to the phosphorylation of ADP.
Subsequent experiments showed that respiration proceeds rapidly until all the ADP has
been phosphorylated (Figure 14.3a) and that the amount of 0 2 consumed depends on
the amount of ADP added.

Synthetic compounds called uncouplers stimulate the oxidation of substrates in the
absence of ADP (Figure 14.3b). The phenomenon of uncoupling helped show how
oxidation reactions are linked to ATP formation. In the presence of an uncoupler, oxygen
uptake (respiration) proceeds until all the available oxygen is consumed. This rapid
oxidation of substrates proceeds with little or no phosphorylation of ADP. In other words,
these synthetic compounds uncouple oxidation from phosphorylation. There are many

14.3 The Chemiosmotic Theory and Protonmotive Force

421

different kinds of uncouplers and they have little in common chemically except that all
are lipid-soluble weak acids. Both their protonated and conjugate base forms can cross
the inner mitochondrial membrane — the anionic conjugate base retains lipid solubility
because the negative charge is delocalized. The resonance structures of the uncoupler
2,4-dmitrophenol are shown in Figure 14.4.

The effect of uncouplers, and many other experiments, revealed that electron
transport (oxygen uptake) and ATP synthesis were normally coupled but the underlying
mechanism was unknown. Throughout the 1960s it was commonly believed that there
must be several steps in the electron transport process where the Gibbs free energy
change was sufficient to drive ATP synthesis. This form of coupling was thought to be
analogous to substrate level phosphorylation.

Mitchell proposed that the action of mitochondrial enzyme complexes generates a
proton concentration gradient across the inner mitochondrial membrane. He suggested
that this gradient provides the energy for ADP phosphorylation via an indirect coupling
to electron transport. Mitchell’s ideas accounted for the effect of the lipid-soluble un-
coupling agents — they bind protons in the cytosol, carry them through the inner mem-
brane, and release them in the matrix, thereby dissipating the proton concentration gra-
dient. The proton carriers uncouple electron transport (oxidation) from ATP synthesis
because protons enter the matrix without passing through ATP synthase.

ATP synthase activity was first recognized in 1948 as ATPase activity in damaged
mitochondria (i.e., damaged mitochondria catalyze hydrolysis of ATP to ADP and Pj).
Most workers assumed that mitochondrial ATPase catalyzes the reverse reaction in un-
damaged mitochondria and this assumption proved to be correct. Efraim Racker and
his coworkers isolated and characterized this membrane-bound oligomeric ATPase in the
1960s. The proton driven reversibility of the ATPase reaction was demonstrated by ob-
serving the expulsion of protons on hydrolysis of ATP in mitochondria. Further support
came from experiments with small membrane vesicles where the enzyme was incorpo-
rated into the membrane. When a suitable proton gradient was created across the vesi-
cle membrane, ATP was synthesized from ADP and Pj (Section 14.9).

B. The Protonmotive Force

Protons are translocated into the intermembrane space by the membrane-associated
electron transport complexes and they flow back into the matrix via ATP synthase.
This circular flow forms a circuit that is similar to an electrical circuit. The energy of
the proton concentration gradient, called the protonmotive force, is analogous to the
electromotive force of electrochemistry (Section 10.9A). This analogy is illustrated in
Figure 14.5.

Consider a reaction such as the reduction of molecular oxygen by the reducing
agent XH 2 in an electrochemical cell.

XH 2 + y 2 0 2 X + H 2 0 (14.1)

(a)

Substrate ADP

(b)

Substrate 2,4-Dinitrophenol

▲ Figure 14.3

Oxygen uptake and ATP synthesis in mitochon-
dria. (a) In the presence of excess Pj and
substrate, intact mitochondria consume oxy-
gen rapidly only when ADP is added. Oxygen
uptake ceases when all the ADP has been
phosphorylated. (b) Adding the uncoupler
2,4-dinitrophenol allows oxidation of the
substrate to proceed in the absence of phos-
phorylation of ADP. The arrows indicate the
times at which additions were made to the
solution of suspended mitochondria.

See Box 15.4 for a description of
Racker’s key experiment.

2,4-Dinitrophenolate anion

▲ Figure 14.4

Protonated and conjugate base forms of 2,4-dinitrophenol. The dinitrophenolate anion is resonance stabilized and its negative ionic charge is broadly
distributed over the ring structure of the molecule. Because the negative charge is delocalized, both the acid and base forms of dinitrophenol are
sufficiently hydrophobic to dissolve in the membrane.

422

CHAPTER 14 Electron Transport and ATP Synthesis

(a)

3 e

■ v 2 o.

a®

■VzOy

a Figure 14.5

Electromotive and protonmotive force, (a) In

an electrochemical cell, electrons pass from
the reducing agent XH 2 to the oxidizing agent
0 2 through a wire connecting the two elec-
trodes. The measured electrical potential
between cells is the electromotive force.

(b) When the configuration is reversed
(i.e., the external pathway for electrons is
replaced by an aqueous pathway for pro-
tons), the potential is the protonmotive
force. In mitochondria, protons are translo-
cated across the inner membrane when
electrons are transported within the mem-
brane by the electron transport chain.

Electrons from XH 2 pass along a wire that connects the two electrodes where the oxida-
tion and reduction half- reactions occur. Electrons flow from the electrode where XH 2 is
oxidized

XH 2 X + 2 H© + 2 e e (14.2)

to the electrode where 0 2 is reduced.

y 2 0 2 + 2 H© + 2 e 0 H 2 0 (14.3)

In the electrochemical cell, protons pass freely from one reaction cell to the other
through the solvent in a salt bridge. Electrons move through an external wire because of
a potential difference between the cells. This potential, measured in volts, is the electro-
motive force. The direction of electron flow and the extent of reduction of the oxidizing
agent depend on the difference in free energy between XH 2 and 0 2 that in turn depends
on their respective reduction potentials.

In mitochondria, it is protons — not electrons — that flow through the external
connection, an aqueous circuit connecting the membrane-associated electron trans-
port chain and ATP synthase. This connection is analogous to the wire of the electro-
chemical reaction. The electrons still pass from the reducing agent XH 2 to the oxidizing
agent 0 2 but in this case it is through the membrane-associated electron transport
chain. The free energy of these oxidation-reduction reactions is stored as the proton-
motive force of the proton concentration gradient and is recovered in the phosphory-
lation of ADP.

Recall from Section 9.10 that the Gibbs free energy change for transport of a
charged molecule is

AG transport = 2.30B/?riog-^4+ zFA'P (14.4)

P%UtJ

where the first term is the Gibbs free energy due to the concentration gradient and the
second term IzFAT^ is due to the charge difference across the membrane. For protons
the charge per molecule is 1 (z = 1.0) and the overall Gibbs free energy change of the
proton gradient is

AG = 2.303 RT

[H©J

'° 9 [H©out]

+ FAT' = 2.303 RT ApH + FAT'

(14.5)

This equation can be used to calculate the protonmotive force generated by the proton
gradient and the charge difference across the membrane. In liver mitochondria the
membrane potential (AT') is —0.17 V (inside negative, Section 9.10A) and the pH dif-
ference is —0.5 (ApH = pH out — pHj n ). The membrane potential is favorable for move-
ment of protons into the mitochondrial matrix so the FAT' term will be negative be-
cause AT' is negative. The pH gradient is also favorable so the first term in Equation 14.5
must be negative. Thus, the equation for protonmotive force is

AG in = FAT' + 2.303 RT ApH

(14.6)

Using the above values at 37° (T = 310 K) the available Gibbs free energy is

AG = [96485 X -0.17] + [2.303 X 8.315 X 310 X -0.5]
= -16402 J mol -1 - 2968 j mol -1 = -19.4 kj mol -1

(14.7)

This means that the transport of a single mole of protons back across the membrane is
associated with a free energy change of — 19.4 kj. That’s a lot of energy for moving such
a small ion!

14.4 Electron Transport 423

The standard Gibbs free energy change for the synthesis of one molecule of ATP
from ADP is 32 kj mol -1 (AG°' = 32 kj mol -1 ) but the actual Gibbs free energy change is
about -48 kj mol -1 (Section 10.6). At least three protons must be translocated in order
to drive synthesis of one ATP molecule (3 x — 19.4 = —58.2 kj mol -1 ) .

Note that 85% ( — 16.4/ — 19.4 = 85%) of the Gibbs free energy change is due to the
charge gradient across the membrane and only 15% ( — 3.0/— 19.4 = 15%) is due to the
proton concentration gradient. Keep in mind that the energy required to create the proton
gradient is +19.4 kj mol -1 .

KEY CONCEPT

The protonmotive force is due to the
combined effect of a charge difference
and a proton concentration difference
across the membrane.

14.4 Electron Transport

We now consider the individual reactions of the membrane- associated electron trans-
port chain. Four oligomeric assemblies of proteins are found in the inner membrane of
mitochondria or the plasma membrane of bacteria. These enzyme complexes have been
isolated in their active forms by careful solubilization using detergents. Each complex
catalyzes a separate portion of the energy transduction process. The numbers I through
IV are assigned to these complexes. Complex V is ATP synthase.

A. Complexes I Through IV

The four enzyme complexes contain a wide variety of oxidation-reduction centers.
These may be co factors such as FAD, FMN, or ubiquinone (Q). Other centers include
Fe-S clusters, heme-containing cytochromes, and copper proteins. Electron flow occurs
via the sequential reduction and oxidation of these redox centers with flow proceeding
from a reducing agent to an oxidizing agent. There are many reactions that involve elec-
tron transport processes in biochemistry. We have already seen several of these reactions
in previous chapters — the flow of electrons in the pyruvate dehydrogenase complex is a
good example (Section 13.1).

Electrons flow through the components of an electron transport chain in the di-
rection of increasing reduction potential. The reduction potentials of each redox
center fall between that of the strong reducing agent, NADH, and that of the termi-
nal oxidizing agent, 0 2 . The mobile coenzymes ubiquinone (Q) and cytochrome c
serve as links between different complexes of the electron transport chain. Q trans-
fers electrons from complexes I or II to complex III. Cytochrome c transfers
electrons from complex III to complex IV. Complex IV uses the electrons for the re-
duction of 0 2 to water.

The order of the electron transport reactions is shown in Figure 14.6 against a scale
of standard reduction potential on the left and a relative scale of Gibbs standard free en-
ergy change on the right. Recall from Section 10.9 that the standard reduction potential
(in units of volts) is directly related to the standard Gibbs free energy change (in units
of kj mol -1 ) by the formula

AG°' = -n F A P'

(14.8)

As you can see from Figure 14.6, a substantial amount of energy is released during the
electron transport process. Much of this energy is stored in the protonmotive force that
drives ATP synthesis. It is this coupling of electron transport to the generation of a pro-
tonmotive force that distinguishes membrane- associated electron transport from other
examples of electron transport.

The values shown in Figure 14.6 are strictly true only under standard conditions
where the temperature is 25°C, the pH is 7.0, and the concentrations of reactants and
products are equal (1M each). The relationship between actual reduction potentials (E)
and standard ones (E or ) is similar to the relationship between actual and standard free
energy (Section 1.4B),

E = E or

RT |n [S red ]

^ [Sox]

= E°'

2.303R7-, [S red ]

(14.9)

The Gibbs Free Energy of Electron
Transport

E = ^acceptor — ^donor (10.26)

_ r°' r°'

~ l 0 2 ~ ^ NADH

= +0.82 - (-0.32) (Table 10.4)
= 1.14 V
A G° = —n?AE°

= -2(96485X1.14)

= 220 kJ mol -1

424 CHAPTER 14 Electron Transport and ATP Synthesis

Cofactors in electron transport

NADH

Succinate

FMN

FAD

Fe-S -
Fe-S -

Fe-S
Cyt b

Cyt c | > Cyt c — >Cyt Cyt a 3 ^ 0 2

0.4

0 . 2 -

0 -

0 . 2 -

0.4-

0 . 6 -

0 . 8 -

NADH -
NAD©^

Complex I
NADH-ubiquinone
oxidoreductase

Succinate

Fumarate

Path of
electron
flow

Complex III

Ubiquinol-cytochrome

oxidoreductase

Complex II

Succinate-ubiquinone

oxidoreductase

(§B>

Complex IV
Cytochrome c
oxidase

r 220

- 165

- 110

- y 2 0 2 + 2 H

^h 2 o

- 55

I- 0

1 . 0-1

▲ Figure 14.6

Electron transport. Each of the four complexes of the electron transport chain, composed of several protein subunits and cofactors, undergoes cyclic
reduction and oxidation. The complexes are linked by the mobile carriers ubiquinone (Q) and cytochrome c. The height of each complex indicates the
A E°' between its reducing agent (substrate) and its oxidizing agent (which becomes the reduced product). Standard reduction potentials are plotted
with the lowest value at the top pf the graph (see Section 10. 9B).

KEY CONCEPT

Aerobic organisms need oxygen because
it serves as the terminal electron
acceptor in membrane-associated
electron transport.

where [S re< j] and [S ox ] represent the actual concentrations of the two oxidation states
of the electron carrier. Under standard conditions, the concentrations of reduced and
oxidized carrier molecules are equal; thus, the ratio [S re< j]/[S ox ] is one, and the second
term in Equation 14.9 is zero. In this case, the actual reduction potential is equal to
the standard reduction potential (at 25°C and pH 7.0). In order for electron carriers
to be efficiently reduced and reoxidized in a linear fashion, appreciable quantities of
both the reduced and oxidized forms of the carriers must be present under steady
state conditions. This is the situation found in mitochondria. We can therefore as-
sume that for any given oxidation-reduction reaction in the electron transport com-
plexes the concentrations of the two oxidation states of the electron carriers are fairly
similar. Since physiological pH is close to 7 under most circumstances and since most
electron transport processes operate at temperatures close to 25°C, we can safely as-
sume that E is not much different from E°' . From now on, our discussion refers only
to E or values.

The standard reduction potentials of the substrates and cofactors of the electron
transport chain are listed in Table 14.1. Note that the values progress from negative to
positive so that, in general, each substrate or intermediate is oxidized by a cofactor or
substrate that has a more positive E or . In fact, one consideration in determining the ac-
tual sequence of the electron carriers was their reduction potentials.

14.4 Electron Transport 425

The Gibbs standard free energy available from the reactions catalyzed by each
complex is shown in Table 14.2. The overall free energy totals -220 kj mol -1 as
shown in Figure 14.6. Complexes I, III, and IV translocate protons across the mem-
brane as electrons pass through the complex. Complex II, which is also the succinate
dehydrogenase complex we examined as a component of the citric acid cycle, does
not directly contribute to formation of the proton concentration gradient. Complex
II transfers electrons from succinate to Q and thus represents a tributary of the res-
piratory chain.

B. Cofactors in Electron Transport

As shown at the top of Figure 14.6, the electrons that flow through complexes I through
IV are actually transferred between coupled cofactors. Electrons enter the membrane-
associated electron transport chain two at a time from the reduced substrates NADH
and succinate. The flavin coenzymes FMN and FAD are reduced in complexes I and II,
respectively. The reduced coenzymes FMNH 2 and FADH 2 donate one electron at a time
and all subsequent steps in the electron transport chain proceed by single electron
transfers. Iron-sulfur (Fe-S) clusters of both the [2 Fe-2 S] and [4 Fe-4 S] type are
present in complexes I, II, and III. Each iron-sulfur cluster can accept or donate one
electron as an iron atom undergoes reduction and oxidation between the ferric [Fe®,
Fe(III)] and ferrous [Fe®, Fe(II)] states. Copper ions and cytochromes are also single
electron oxidation-reduction agents.

Several different cytochromes are present in the mammalian mitochondrial en-
zyme complexes. These include cytochrome fr L , cytochrome fr H , cytochrome q, cy-
tochrome a, and cytochrome a 3 . Very similar cytochromes are found in other species.
Cytochromes transfer electrons from a reducing agent to an oxidizing agent by cy-
cling between the ferric and ferrous oxidation states of the iron atoms of their heme
prosthetic groups (Section 7.17). Individual cytochromes have different reduction
potentials because of differences in the structures of their apoproteins and sometimes
their heme groups (Table 14.1). These differences allow heme groups to function as
electron carriers at several points in the electron transport chain. Similarly, the reduc-
tion potentials of iron-sulfur clusters can vary widely depending on the local protein
environment.

The membrane- associated electron transport complexes are functionally linked by
the mobile electron carriers ubiquinone (Q) and cytochrome c. Q is a lipid-soluble
molecule that can accept and donate two electrons, one at a time (Section 7.15). Q dif-
fuses within the lipid bilayer accepting electrons from complexes I and II and passing
them to complex III. The other mobile electron carrier is cytochrome c, a peripheral
membrane protein associated with the outer face of the membrane. Cytochrome c
carries electrons from complex III to complex IV. The structures and the oxidation-
reduction reactions of each of the four electron transport complexes are examined in
detail in the following sections.

Table 14.1 Standard reduction potentials
of mitochondrial oxidation-
reduction components

Substrate
of Complex

E°' (V)

NADH

-0.32

Complex 1

FMN

-0.30

Fe-S clusters

-0.25 to -0.05

Succinate

+0.03

Complex II

FAD

0.0

Fe-S clusters

-0.26 to 0.00

QH 2 /Q

+0.04

(•Q e /Q

-0.16)

(QH 2 /-Q©

+0.28)

Complex III

Cytochrome b \.

- 0.01

Cytochrome b H

+0.03

Fe-S cluster

+0.28

Cytochrome C|

+0.22

Cytochrome c

+0.22

Complex IV

Cytochrome a

+0.21

Cu A

+0.24

Cytochrome 03

+0.39

Cu B

+0.34

o 2

+0.82

KEY CONCEPT

The transfer of electrons from NADH to 0 2
releases enough energy to drive synthesis
of many ATP molecules.

Table 14.2 Standard free energy released in the oxidation reaction catalyzed by each complex

Complex

r°'

c reductant

(V)

r°'

c oxidant

(V)

AE°'°

(V)

AC°' b

(kj mol 1 )

1 (NADH/Q)

-0.32

-0.04

+0.36

-60

II (Succinate/Q)

+0.03

+0.04

+0.01

-2

III (QH 2 /Cytochrome c)

+0.04

+0.22

+0.18

-35

IV (Cytochrome c/0 2 )

+0.22

+0.82

+0.59

-116

°A E°' was calculated as the difference between P/eductant and ^oxidant.

^The Gibbs standard free energy was calculated using Equation 14.8 where n = 2 electrons.

426 CHAPTER 14 Electron Transport and ATP Synthesis

▲ Figure 14.7

Structure of complex I. The structures of com-
plex I have been determined at low resolution
by analyzing electron micrographic images,
(a) Complex I from the bacterium Aquifex
aeolicus. (b) Complex I from cow, Bos taurus.
(c) Complex I from the yeast, Yarrowia
lipolytica.

Outer membrane Inner membrane

14.5 Complex I

Complex I catalyzes the transfer of two electrons from NADH to Q. The systematic
name of this enzyme is NADH:ubiquinone oxidoreductase. It is a very complicated en-
zyme whose structure has not been completely solved. The prokaryotic versions contain
14 different polypeptide chains. The eukaryotic forms have 14 homologous subunits
plus 20-32 additional subunits, depending on the species. The extra eukaryotic sub-
units probably stabilize the complex and prevent electron leakage.

The structure of the complex is L-shaped as seen in the electron microscope
(Figure 14.7). The membrane-bound component consists of multiple subunits that
span the membrane. This module contains a proton transporter activity. A larger
component projects into the mitochondrial matrix, or the cytoplasm in bacteria
(Figure 14.8). This arm contains a terminal NADH dehydrogenase activity and FMN.
The connector module is composed of multiple subunits with 8 or 9 Fe-S clusters
(Figure 14.9).

NADH molecules on the inside surface of the membrane donate electrons to
complex I. The electrons are passed two at a time as a hydride ion (H®, two electrons
and a proton). In the first step of electron transfer the hydride ion is transferred
to FMN forming FMNH 2 . FMNH 2 is then oxidized in two steps via a semiquinone
intermediate. The two electrons are transferred one at a time to the next oxidizing
agent, an iron-sulfur cluster.

+ H©. + H©

FMN > FMNH 2

-H©, -e©

» FMNH

-H©, -e©

* FMN

(14.10)

FMN is a transducer that converts two-electron transfer from NAD-linked dehy-
drogenases to one-electron transfer for the rest of the electron transport chain. In
complex I the cofactor FMNH 2 transfers electrons to sequentially linked iron-sulfur
clusters. There are at least eight Fe-S clusters positioned within the same arm of com-
plex I that contains the NADH dehydrogenase activity. These Fe-S clusters provide a
channel for electrons by directing them to the membrane-bound portion of the com-
plex where ubiquinone (Q) accepts electrons one at a time passing through a
semiquinone anion intermediate (*Q®) before reaching its fully reduced state,
ubiquinol (QH 2 ).

+ e©

•Q©

+ e©, +2 H©

* QH 2

(14.11)

Q and QH 2 are lipid-soluble cofactors. They remain within the lipid bilayer and
can diffuse freely in two dimensions. Note that the Q binding site of complex I is within
the membrane. One of the reasons for the complicated electron transport chain within
complex I is to carry electrons from an aqueous environment to a hydrophobic envi-
ronment within the membrane.

As electrons move through complex I, two protons (one originating from the hy-
dride ion of NADH and one from the interior) are transferred to FMN to form
FMNH 2 . These two protons or their equivalents are consumed in the reduction of Q to
QH 2 . Thus, two protons are taken up from the interior and transferred to QH 2 . They
are not released to the exterior in the complex I reactions. (QH 2 is subsequently reoxi-
dized by complex III and the protons are then released to the exterior. This is part of the
proton translocation activity of complex III described in Section 14.7.)

In complex I, four protons are directly translocated across the membrane for every
pair of electrons that pass from NADH to QH 2 . These do not include the protons re-
quired for ubiquinone reduction. The proton pump is probably an H©/Na© antiporter

◄ Figure 14.8

Complex orientation. The electron transport complexes are embedded in the inner membrane. They
can be drawn with the outside of the membrane at the top or at the bottom of the figure. Both
views are seen in the scientific literature. We have chosen the orientation with the outside on top
and the inside of the matrix on the bottom.

14.6 Complex II 427

Connector
module 2 H 0

2H©

Transporter module 4 H 0

OUTSIDE

Fe-S

4 H

INSIDE

NADH + H

NADH dehydrogenase
2e © module

NAD 0

◄ Figure 14.9

Electron transfer and proton flow in Complex I.

Electrons are passed from NADH to Q via
FMN and a series of Fe-S clusters. The re-
duction of Q to QH 2 requires two protons
taken up from the inside compartment.

In addition, four protons are translocated
across the membrane for each pair of
electrons transferred.

located in the membrane-bound module. The mechanism of proton translocation is
not clear — it is likely coupled to conformational changes in the structure of complex I
as electrons flow from the NADH dehydrogenase site to the ubiquinone binding site.

Heme b

14.6 Complex II

Complex II is succinate:ubiquinone oxidoreductase, also called the succinate dehydroge-
nase complex. This is the same enzyme that we encountered in the previous chapter
(Section 13.3#6). It catalyzes one of the reactions of the citric acid cycle. Complex II ac-
cepts electrons from succinate and, like complex I, catalyzes the reduction of Q to QH 2 .

Complex II contains three identical multisubunit enzymes that associate to form a
trimeric structure that is firmly embedded in the membrane (Figure 14.10). The over-
all shape resembles a mushroom with its head projecting into the interior of the
membrane compartment. Each of the three succinate dehydrogenase enzymes has two
subunits forming the head and one or two subunits (depending on the species)
forming the membrane-bound stalk. One of the head subunits contains the
substrate binding site and a covalently bound flavin adenine dinucleotide
(FAD). The other head subunit contains three Fe-S clusters.

The head subunits from all species are closely related and share significant
sequence similarity with other members of the succinate dehydrogenase family
(e.g., fumarate reductase, Section 14.13). The membrane subunits, on the other
hand, may be very different (and unrelated) in various species. In general, the
membrane component has one or two subunits that consist exclusively of
membrane-spanning a helices. Most of them have a bound heme b molecule
and this subunit is often called cytochrome b. All of the membrane subunits
have a Q binding site positioned near the interior surface of the membrane at
the point where the head subunits are in contact with the membrane subunits.

The sequence of reactions for the transfer of two electrons from succinate
to Q begins with the reduction of FAD by a hydride ion. This is followed by two
single electron transfers from the reduced flavin to the series of three iron-
sulfur clusters (Figure 14.1 1). (In those species with a cytochrome b anchor, the
heme group is not part of the electron transfer pathway.)

Very little free energy is released in the reactions catalyzed by complex II
(Table 14.2). This means that the complex cannot contribute directly to the
proton concentration gradient across the membrane. Instead, it supplies electrons
from the oxidation of succinate midway along the electron transport sequence.

Q can accept electrons from complex I or II and donate them to complex III
and then to the rest of the electron transport chain. Reactions in several other
pathways also donate electrons to Q. We saw one of them, the reaction cat-
alyzed by the glycerol 3-phosphate dehydrogenase complex, in Section 12. 2C.

OUTSIDE

-Membrane

INSIDE

Fe-S clusters

FAD

▲ Figure 14.10

Structure of the E. coli succinate dehydrogenase complex.

A single copy of the enzyme showing the positions
of FAD, the three Fe-S clusters, QH 2 , and heme b.
Complex II contains three copies of this multisubunit
enzyme. [PDB 1NEK]

428 CHAPTER 14 Electron Transport and ATP Synthesis

Figure 14.1 1 ►

Electron transfer in complex II. A pair of elec-
trons is passed from succinate to FAD as
part of the citric acid cycle. Electrons are
transferred one at a time from FADH 2 to
three Fe-S clusters and then to Q. (Only
one Fe-S cluster is shown in the figure.)
Complex II does not directly contribute to
the proton concentration gradient but serves
as a tributary that supplies electrons (as
QH 2 ) to the rest of the electron transport
chain.

Complex III is, arguably, the most
important enzyme in metabolism.

Very similar complexes are present in
chloroplasts where they participate in
electron transport and proton translo-
cation during photosynthesis.

14.7 Complex III

Complex III is ubiquinol: cytochrome c oxidoreductase, also called the cytochrome frq
complex. This enzyme catalyzes the oxidation of ubiquinol (QH 2 ) molecules in the
membrane and the reduction of a mobile water-soluble cytochrome c molecule on the
exterior surface. Electron transport through complex III is coupled to the transfer of
H© across the membrane by a process known as the Q cycle.

The structures of the cytochrome frq complexes from many bacterial and eukaryotic
species have been solved by X-ray crystallography. Complex III contains two copies of
the enzyme and is firmly anchored to the membrane by a large number of a helices that
span the lipid bilayer (Figure 14.12). The functional enzyme consists of three main subunits:
cytochrome q, cytochrome b , and the Rieske iron sulfur protein (ISP) (Figure 14.13).
(Note that the cytochrome q subunit is a different protein than the mobile cytochrome
c product of the reaction.) Other subunits are present on the inside surface but they do
not play a direct role in the ubiquinokcytochrome c oxidoreductase reaction. The mo-
bile cytochrome c electron acceptor binds at the top of the complex — the part that is ex-
posed to the exterior side of the membrane.

Figure 14.12 ►

Complex III from cow ( Bos taurus) mitochon-
dria. The cytochrome bc\ complex contains
two copies of the enzyme ubiquinone:
cytochrome c oxidoreductase. [PDB 1PP9]

14.7 Complex III 429

The path of electrons through the complex is shown in Figure 14.14. The reaction
begins when QH 2 (from complex I or complex II) binds to the Q 0 site in the cy-
tochrome b subunit. QH 2 is oxidized to the semiquinone and a single electron is passed
to the adjacent Fe-S complex in the ISP subunit. From there, the electron transfers to
the heme group in cytochrome q. This transfer is facilitated by movement of the head
group of ISP. In the electron accepting position, the Fe-S cluster is adjacent to the Q 0
site and in the electron donating position the Fe-S cluster shifts to a position near the
heme group in cytochrome q. Soluble cytochrome c is oxidized by transfer of an elec-
tron from the membrane-bound cytochrome q subunit of complex III.

In this reaction, the terminal electron acceptor is cytochrome c (Section 7.17). This
molecule serves as a mobile electron carrier transferring electrons to complex IV, the
next component of the chain (Figure 14.15). The role of reduced cytochrome c is simi-
lar to that of QH 2 , which carries electrons from complex I to complex III. The structures
of cytochrome c electron carriers from all species are remarkably similar (Section 4.7B,
Figure 4.21) and the amino acid sequences of the polypeptide chain are well conserved
(Section 3.11, Figure 3.23).

The oxidation of QH 2 at the Q 0 site is a two-step process with a single electron
transferred at each step. The path of electrons from the second step, oxidation of the
semiquinone intermediate, follows a different route than the first electron. In this case,
the electron is passed sequentially to two different fr-type hemes within the membrane
portion of the complex. The first heme group (fr L ) has a lower reduction potential and
the second heme (fr H ) has a higher reduction potential (Table 14.1).

The b H heme is part of the Q : site where a molecule of Q is reduced to QH 2 in a
two-step reaction that involves a semiquinone intermediate. A single electron is trans-
ported from b L (at the Q 0 site) to b H (at the Q x site) to Q to produce the semiquinone.

▲ Figure 14.13

Subunits of complex III. The three catalytic
subunits of each dimer are Cytochrome q
(green), cytochrome b (blue) and the Rieske
iron sulfur protein (ISP) (red). Cytochrome c
(dark blue) binds to the Cytochrome q sub-
unit. [PDB 1PP9]

Cytochrome C (x2)

2 hr

Cytochrome b

▲ Figure 14.14

Electron transfer and proton flow in complex III. Two pairs of electrons are passed separately from two
molecules of QH 2 at the Q 0 site. Each pair of electrons is split so that individual electrons follow
separate pathways. One electron is transferred to an Fe-S cluster then to cytochrome q and finally
to cytochrome c, the terminal electron carrier. The other electron from each oxidation of QH 2 is
transferred to heme b H (Qi site) and then to Q. A total of four protons are translocated across the
membrane: two from the inside compartment and two from QH 2 . (Only the left-hand half of the
dimer is shown and the bottom subunits that project into the matrix are not shown.)

430 CHAPTER 14 Electron Transport and ATP Synthesis

Oxidized

Reduced

▲ Figure 14.15

Cytochrome c. Oxidized (top) and reduced
(bottom) forms of cytochrome c from horse
( Equus cabal I us). The iron atom in the cen-
ter of the heme group (orange) shifts from
Fe© to Fe© as it gains an electron from
complex III. This reduction is accompanied
by small changes in the conformation of the
protein. [PDB 10CD (top) 1GIW (bottom)]

Table 14.3

Qo : 2QH 2 + 2cytc(Fe®) ► 2 Q + 2 cyt c(Fe®) + 2 e 0 + 4 H© out

Qi : Q + 2 H© in + 2 e© * QH 2

Sum: QH 2 + 2 cyt c(Fe®) + 2 H© in Q + 2 cyt c(Fe®) + 4 H© out

Then, a second electron is transferred to reduce the semiquinone to QH 2 . The second
electron is derived from the oxidation of a second molecule of QH 2 at the Q 0 site. This
second oxidation of QH 2 also results in the reduction of a second molecule of cy-
tochrome c since the two electrons from the second QH 2 follow separate paths. The net
result is that the oxidation of two molecules of QH 2 at the Q 0 site produces two mole-
cules of reduced cytochrome c and regenerates a molecule of QH 2 at the site. The
two cycles of QH 2 oxidation are shown in Figure 14.16. The entire pathway is known as
the Q cycle and it is one of the most important reactions in all of metabolism because it
is the one most responsible for creating the protonmotive force.

Four protons are produced during the oxidation of two molecules of QH 2 at the Q 0
site. These protons are released to the exterior of the membrane compartment and they
contribute to the proton gradient that is formed during membrane-associated electron
transport. The protons originate in the interior compartment. They may have been
taken up in the reactions catalyzed by complex I or complex II or they may be derived
from protons taken up on the inside of the membrane during reduction of Q at the
site in complex III as shown in Figure 14.16.

The stoichiometry of the complete Q-cycle reaction is shown in Table 14.3. For
every pair of electrons that pass through complex III from QH 2 to cytochrome c there
are four protons translocated across the membrane. Two molecules of cytochrome c are
reduced and these mobile carriers transport one electron each to complex IV. Note that
there are actually two molecules of QH 2 oxidized (giving up four electrons) but two of
these electrons are recycled to regenerate a molecule of QH 2 .

The complete reaction catalyzed by ubiquinoneicytochrome c oxidoreductase
(complex III) includes the Q cycle and proton translocation across the membrane.
The complex III reaction is a fine example of the relationship between structure and
function. While the stoichiometry of the Q cycle had been known for many years, the
actual mechanism of the reaction only became apparent once the complete structure
was solved in 1998.

KEY CONCEPT

The net effect of the Q cycle is transfer of
four protons to the exterior of the membrane
for every two electrons transferred from
QH 2 to cytochrome c.

Cytochrome c

Q. Q
Cycle 1

Cytochrome c

Figure 14.16 ►

Q cycle. A molecule of QH 2 is oxidized in cycle 1 and a separate molecule is oxidized in cycle 2. Each
cycle produces a molecule of reduced cytochrome c. The combination of cycle 1 and cycle 2 results in
a two-stage reduction of Q to QH 2 . Four protons are released on the exterior side of the membrane.

14.8 Complex IV

431

14.8 Complex IV

Complex IV is cytochrome c oxidase. This complex catalyzes the oxidation of the re-
duced cytochrome c molecules produced by complex III. The reaction includes a four-
electron reduction of molecular oxygen (0 2 ) to water (2 H 2 0) and translocation of
four protons across the membrane.

Complex IV contains two functional units of cytochrome c oxidase. Each cy-
tochrome c oxidase contains single copies of subunits I, II, and III (Figure 14.17). The
bacterial enzymes contain only one additional subunit in each functional unit but the
eukaryotic (mitochondrial) enzymes have up to ten additional subunits. Additional
subunits in the eukaryotic complexes play a role in assembling complex IV and in stabi-
lizing the structure.

The core structure of cytochrome c oxidase is formed from the three conserved
subunits — I, II, and III. These polypeptides are encoded by mitochondrial genes in all
eukaryotes. Subunit I is almost entirely embedded in the membrane. The bulk of this
polypeptide consists of 12 transmembrane a helices. There are three redox centers
buried within subunit I — two of them are a- type hemes (heme a and heme a 3 ) and the
third is a copper ion (Cu B ). The copper atom is in close proximity to the iron atom of
heme a 3 forming a binuclear center where the reduction of molecular oxygen takes
place (Figure 14.18).

Subunit II has two transmembrane helices that anchor it to the membrane. Most
of the polypeptide chain forms a /3-barrel domain located on the exterior surface of
the membrane. This domain contains a copper redox center (Cu A ) composed of two
copper ions. These two copper atoms share electrons forming a mixed valence state.
The external domain of subunit II is the site where cytochrome c binds to cytochrome c
oxidase.

Subunit III has seven transmembrane helices and is completely embedded in the
membrane. There are no redox centers in subunit III and it can be artificially removed
without loss of catalytic activity. Its role in vivo is to stabilize subunits I and II and help
protect the redox centers from inappropriate oxidation-reduction reactions.

Figure 14.19 shows the sequence of electron transfers in complex IV. Cytochrome c
binds to subunit II and transfers an electron to the Cu A site. The pair of copper ions at
the Cu A site can accept and donate one electron at a time — much like an Fe-S cluster.
The complete oxidation of 0 2 requires four electrons. Thus, four cytochrome c molecules
have to bind and sequentially transfer a single electron each to the Cu A redox center.

▲ Figure 14.18

Redox centers in cytochrome c oxidase.

Organization of the heme and copper cofac-
tors in one of the cytochrome c oxidase units.
[PDB 10CC]

◄ Figure 14.17

Structure of cow ( Bos taurus ) complex IV from
mitochondria. The complex consists of two
functional units of cytochrome c oxidase.
Each unit is composed of 13 subunits with
multiple membrane-spanning a helices.
[PDB 10CC]

432 CHAPTER 14 Electron Transport and ATP Synthesis

Figure 14.19 ►

Electron transfer and proton flow in complex IV.

The iron atoms of the heme groups in the a
cytochromes and the copper atoms are both
oxidized and reduced as electrons flow from
cytochrome cto oxygen. Electron transport
through complex IV is coupled to the trans-
fer of protons across the membrane. The
diagram shows the stoichiometry for transfer
of a pair of electrons as in previous figures.
The actual reaction involves the transfer of
four electrons to a molecule of 0 2 to form
two molecules of water.

Cytochrome c (¥2)

2 H® '/ 2 O 2 + 2 H© H 2 0

▲ Figure 14.20

Mitochondrial genome. Mitochondrial genomes
are small, circular, double-stranded DNA
molecules. They contain genes for ribosomal
RNAs (12S rRNA, 16S rRNA) and tRNAs
(labeled according to the amino acid they
carry). The human mitochondrial genome,
shown here, is only 16,589 bp in size and it
encodes only a few of the subunits of the
electron transport complexes. Genes for the
subunits of complex I are colored green, a
complex III subunit is purple, complex IV
subunits are pink, and complex V subunits
are yellow. The D-loop is a highly variable
region required for DNA replication.
Sequences of individual D-loop regions have
been used to trace the evolution of modern
humans providing early evidence that we all
descend from a population in Africa.

Electrons are transferred one at a time from the Cu A site to the heme a prosthetic
group in subunit I. From there they are transferred to the heme a 3 -Cu B binuclear cen-
ter. The two heme groups (a and a 3 ) have identical structures but differ in their stan-
dard reduction potentials due to the local microenvironment formed by surrounding
amino acid side chains in subunit I. Electrons can accumulate at the binuclear center as
the heme iron alternates between Fe© and Fe© states and the copper atom shifts
from Cu© to Cu®. The detailed mechanism for reduction of molecular oxygen at the
binuclear center is under active investigation in a number of laboratories. The first step
involves the rapid splitting of molecular oxygen. One oxygen atom is bound to the iron
atom of the a 3 -heme group and the other is bound to the copper atom. Subsequent
protonation and electron transfer results in the release of a water molecule from the
copper site followed by release of a second water molecule from the iron ligand. The
overall reaction requires the uptake of four protons from the inside surface of the
membrane

0 2 + 4 e© + 4 H© in * 2 H 2 0 (14.12)

The site where oxygen is reduced is buried within the protein in the middle of the lipid
bilayer of the membrane. Charged protons cannot access this site by passive diffusion —
instead, the enzyme contains a channel leading from the inside of the membrane to the
active site. This channel is filled with a single line of water molecules that rapidly ex-
change protons leading to the net movement of protons along this “water wire.”

The reactions of cytochrome c oxidase are coupled to the transfer of protons across
the membrane. One proton is translocated for each electron that passes from
cytochrome c to the final product (H 2 0). The protons move through a water- filled
channel in complex IV and this movement is driven by conformational changes in the
enzyme as oxygen is reduced. The stoichiometry of the complete reaction catalyzed by
complex IV is

4 cyt P + 0 2 + 8 H© in > 4 cyt c @ + 2 H 2 0 + 4 H© out (14.13)

Complex IV contributes to the proton gradient that will drive ATP synthesis. Two
protons are translocated for each pair of electrons that pass through this complex.
Recall that complex I transfers four protons for each pair of electrons and complex III
also translocates four protons for each electron pair. Thus, the membrane-associated
electron transport system pumps ten protons across the membrane for every molecule
of NADH that is oxidized.

The genes encoding the various subunits of the mitochondrial complexes may
be in the nucleus or the mitochondria, depending on the species. The genes for
cytochrome c oxidase subunits are always found in the mitochondrial genome
(Figure 14.20).

14.9 Complex V: ATP Synthase 433

14.9 Complex V: ATP Synthase

Complex V is ATP synthase. It catalyzes the synthesis of ATP from ADP + Pj in a reac-
tion that is driven by the proton gradient generated during membrane-associated elec-
tron transport. ATP synthase is a specific F-type ATPase called FqF! ATPase — named
after the reverse reaction. In spite of its name, F-type ATPase is responsible for
synthesizing ATP — not hydrolyzing it. The enzyme is embedded in the membrane and
has a characteristic knob -and- stalk structure that has been observed in electron micro-
graphs for over half a century (Figure 14.21). The F x (knob) component contains the
catalytic subunits — when released from membrane preparations it catalyzes the hydrol-
ysis of ATP. For this reason, it has traditionally been referred to as F x ATPase. This part
of the enzyme projects into the mitochondrial matrix in eukaryotes and into the cyto-
plasm in bacteria. (ATP synthase is also found in chloroplast membranes, as we will see
in the next chapter.) The F 0 (stalk) component is embedded in the membrane. It has a
proton channel that spans the membrane, and the passage of protons through this
channel from the outside of the membrane to the inside is coupled to the formation of
ATP by the F x component.

Recent cryo electron micrograph structures of ATP synthase have revealed details
of its overall structure. These can be correlated with the X-ray crystallographic struc-
tures of the various components (Figure 14.22).

The subunit composition of the F x component (knob) is a 3 /3 3 y8s and that of the
F 0 membrane component is a 1 b 2 c 10 _ 14 . The c subunits of F 0 interact to form a cylindri-
cal base within the membrane. The core of the F x (knob) structure is formed from three
copies each of subunits a and /3 arranged as a cylindrical hexamer. The nucleotide bind-
ing sites lie in the clefts between adjacent a and (3 subunits. Thus, the binding sites are
spaced 120° apart on the surface of the a 3 /? 3 cylinder. The catalytic site of ATP synthesis
is mostly associated with amino acid residues of the (3 subunit.

▲ Figure 14.21

Knobs and stalks. The internal mitochondrial
membranes are studded with structures that
look like knobs projecting into the mitochon-
drial matrix at the end of short membrane-
embedded stalks.

F 0

“303

C 1 0-1 5

Periplasm

Cytoplasm

◄ Figure 14.22

ATP synthase structure. The Fi component is
on the inner face of the membrane. The F 0
component, which spans the membrane, forms
a proton channel at the interface between the
a and c subunits. The passage of protons
through this channel causes the c subunit
rotor (blue) to rotate relative to the stator of
a and b subunits (orange). The torque of these
rotations is transmitted to Fi where it is used
to drive ATP synthesis as the /subunit (cyan)
rotates within the head formed by a and /I sub-
units (green). (The e subunit is part of the stalk —
it lies behind the y subunit in this view.)
(Modified from von Ballmoos et al . , 2009.)

434

CHAPTER 14 Electron Transport and ATP Synthesis

V-ATPases have a similar structure.
They use ATP hydrolysis to drive the
import of protons into acidic vesicles
(vacuoles). This is the reverse of the
reaction catalyzed by ATP synthase.

t Figure 14.23

Binding change mechanism of ATP synthase.

The different conformations of the three cat-
alytic sites are indicated by different shapes.
ADP and Pj bind to the yellow site in the open
conformation. As the /shaft rotates in the
counterclockwise direction (viewed from the
cytoplasmic/matrix end of the Fi component),
the yellow site is converted to a loose confor-
mation where ADP and Pj are more firmly
bound. Following the next step of the rotation
the yellow site is converted to a tight confor-
mation and ATP is synthesized. Meanwhile,
the site that had bound ATP tightly has be-
come an open site and a loose site containing
other molecules of ADP and Pj has become a
tight site. ATP is released from the open site
and ATP is synthesized in the tight site.

The a 3 (5 3 oligomer of F x is connected to the transmembrane c subunits by a multi-
subunit stalk made up of the y and s subunits. The c-s-y unit forms a “rotor” that spins
within the membrane. Rotation of the y subunit inside the a 3 p 3 hexamer alters the
conformation of the /3 subunits, opening and closing the active sites. The a, b, and S
subunits form an arm that also attaches the F 0 component to the a 3 p 3 oligomer. This
a-b-S-a 3 fi 3 unit is termed the “stator” Passage of protons through the channel at the in-
terface between the a and c subunits causes the rotor assembly to spin in one direction
relative to the stator. The entire structure is often called a molecular motor.

There are 10-14 c subunits in the membrane-associated c-ring at the base of the
rotor. The number of subunits depends on the species — yeast and E. coli have a 10-subunit
ring but plants and animals have up to 14 subunits. There is good evidence to indicate
that the rotation of each c subunit past the stator is driven by translocation of a single
proton. Rotation of the /subunit within the F x component takes place in a stepwise,
jerky manner where each step is 120° of rotation. As the c-ring rotates it twists the /
shaft until enough tension builds up to cause it to snap into the next position within the
a 3 p 3 hexamer. If the c-ring has ten subunits then a complete rotation requires translo-
cation of ten protons and results in the production of three ATP molecules but the exact
stoichiometry is still being worked out. The results of many experiments indicate that,
on average, three protons must be translocated for each ATP molecule synthesized and
that’s the value that we will use in the rest of this book. It suggests that only nine proton
translocations are required for one complete rotation of the c-ring.

The mechanism of ATP synthesis from ADP and Pj has been the target of intensive
research for several decades. In 1979 Paul Boyer proposed the binding change mecha-
nism based on observations suggesting that the substrate and product binding proper-
ties of the active site could change as protons moved across the membrane. The a 3 (5 3
oligomer of ATP synthase contains three catalytic sites. At any given time, each site can
be in one of three different conformations: (1) open: newly synthesized ATP can be re-
leased and ADP + Pi can bind; (2) loose: bound ADP + Pi cannot be released; (3) tight:
ATP is very tightly bound and condensation of ADP + Pi is favored. All three sites pass
sequentially through these conformations as the / subunit rotates within the knob. The
rate of this reaction is comparable to that of many enzymes. The rotor turns at ten rev-
olutions per second producing 30 ATP molecules per second. Typical turnover numbers
(fc cat ) are in the range of 100-1000 reactions per second.

The formation and release of ATP are believed to occur by the following steps,
summarized in Figure 14.23:

1. One molecule of ADP and one molecule of Pi bind to an open site.

2. Rotation of the y shaft causes each of the three catalytic sites to change conforma-
tion. The open conformation (containing the newly bound ADP and Pi) becomes a
loose site. The loose site, already filled with ADP and P i? becomes a tight site. The
site containing ATP becomes an open site.

3. ATP is released from the open site and ADP and Pi condense to form ATP in the
tight site.

ATP ADP + P,

14.10 Active Transport of ATP, ADP, and Pj Across the Mitochondrial Membrane 435

BOX 14.2 PROTON LEAKS AND HEAT PRODUCTION

Proton leaks appear to be a major consumer of free energy in
mammals. In a resting adult mammal, about 90% of oxygen
consumption takes place in the mitochondria and about 80%
of this is coupled to ATP synthesis. Quantitative estimates in-
dicate that the ATP produced by mitochondria is used for
protein synthesis (almost 30% of the available ATP), for active
transport of ions by Na© — K© ATPase and Ca© ATPase
(25% to 35%), for gluconeogenesis (up to 10%), and for
other metabolic processes including heat generation. A sig-
nificant amount of the energy from oxidation is not used for
the synthesis of ATP. In resting mammals, at least 20% of the
oxygen consumed by mitochondria is uncoupled by mito-
chondrial proton leakage. This leakage produces heat directly
without apparent use.

The generation of heat in newborns and hibernating an-
imals is a special example of deliberate uncoupling of proton
translocation and ATP synthesis. This physiological uncou-
pling occurs in brown adipose tissue, whose brown color is
due to its many mitochondria. Brown adipose tissue is found
in abundance in newborn mammals and in species that hi-
bernate. The free energy of NADH is not conserved as ATP
but is lost as heat because oxidation is uncoupled from phos-
phorylation. The uncoupling is due to uncoupling protein 1
(UCP1, thermogenin) that forms a channel for the re-entry
of protons into the mitochondrial matrix. When UCP1 is ac-
tive the free energy released is dissipated as heat, raising the
body temperature of the animal.

The strongest evidence that ATP synthase is a rotating motor has been ob-
tained using the a 3 /3 3 y complex immobilized on a glass plate and modified by
attachment of a fluorescent actin filament (Figure 14.24). Rotation of single mol-
ecules was observed by microscopy in the presence of ATP. In this experiment, the
labeled y subunit rotates inside the a 3 /? 3 oligomer in response to ATP hydrolysis.
This rotation is counterclockwise as depicted in Figure 14.24. Note that rota-
tion driven by ATP hydrolysis is in the opposite direction to that observed when
rotation is driven by the proton gradient and ATP is synthesized. The rotation of
the y shaft took place in 120° increments with one step for each ATP molecule hy-
drolyzed. Under ideal conditions, rates of more than 130 revolutions per second
have been observed. This is the expected rotation rate based on the measured
rate of ATP hydrolysis. It is much faster than the in vivo rate of rotation during
ATP synthesis.

14.10 Active Transport of ATP, ADP, and

Pj Across the Mitochondrial Membrane

A large fraction of the total ATP synthesized in eukaryotic cells is made in the
mitochondria. These molecules must be exported since most of them are used in the
cytoplasm. An active transporter is required to allow ADP to enter and ATP to leave mi-
tochondria because the inner mitochondrial membrane is impermeable to charged sub-
stances. This transporter is called the adenine nucleotide translocase — it exchanges
mitochondrial ATP and cytosolic ADP (Figure 14.25). Normally adenine nucleotides
are complexed with Mg© but this is not the case when they are transported across the
membrane. Exchange of ADP© and ATP© causes the loss of a net charge of -1 in the
matrix. This type of exchange draws on the electrical part of the protonmotive force
(AM/ 1 ) and some of the free energy of the proton concentration gradient is expended to
drive this transport process.

The formation of ATP from ADP and Pj in the mitochondrial matrix also requires
a phosphate transporter to import Pi from the cytosol. Phosphate (H 2 P0 4 _ ) is trans-
ported into mitochondria in electroneutral symport with H© (Figure 14.25). The
phosphate carrier does not draw on the electrical component of the protonmotive
force but does draw on the concentration difference, ApH. Thus, both transporters
necessary for ATP formation use up some of the protonmotive force generated by pro-
ton translocation. The combined energy cost of transporting ATP out of the matrix
and ADP and Pi into it is approximately equivalent to the influx of one proton. There-
fore, the synthesis of one molecule of cytoplasmic ATP by ATP synthase requires the

▲ Figure 14.24

Demonstration of the rotation of a single mole-
cule of ATP synthase, complexes were
bound to a glass coverslip and the ysubunit
was attached to a long fluorescent protein
arm. The arms on the molecules rotated
when ATP was added. [Adapted from Noji,
H., Yasuda, R., Yoshida, M., and Kinosita,
K., Jr. (1997). Direct observation of rotation
of Fi -ATPase. Nature 386:299-302.]

KEY CONCEPT

The chemical energy of the protonmotive
force is converted to mechanical energy
by causing the rotation of the ATP
synthase rotor.

Active transport by ATPases is dis-
cussed in Section 9.1 ID.

436 CHAPTER 14 Electron Transport and ATP Synthesis

Figure 14.25 ►

Transport of ATP, ADP, and Pj across the inner
mitochondrial membrane. The adenine nu-
cleotide translocase carries out unidirec-
tional exchange of ATP for ADP (antiport).
Note that the symport of Pj and H® is
electroneutral.

net influx of four protons from the intermembrane space — one for transport and
three that pass through the F 0 component of ATP synthase. Bacteria do not need to
transport ATP or ADP across a membrane so the overall expense of ATP synthesis is
less than that in eukaryotic cells.

KEY CONCEPT

The oxidation of a molecule of NADH
results in the synthesis of 2.5 molecules
of ATP. In terms of metabolic currency,
one NADH molecule is 2.5 ATP
equivalents.

14.11 The P/O Ratio

Before the chemiosmotic theory was proposed, many researchers were searching for a
“high energy” intermediate capable of forming ATP by direct phosphoryl group transfer.
They assumed that complexes I, III, and IV each contributed to ATP formation with
one-to-one stoichiometry. We now know that energy transduction occurs by generating
and consuming a proton concentration gradient. The yield of ATP need not be equiva-
lent for each proton translocating electron transport complex nor must the yield of ATP
per molecule of substrate oxidized be an integral number.

Many different membrane-associated electron transport complexes contribute si-
multaneously to the proton concentration gradient. This common energy reservoir is
drawn on by many ATP synthase complexes. We saw in the preceding sections that the
formation of one molecule of ATP from ADP and P* catalyzed by ATP synthase requires
the inward passage of about three protons and one more proton is needed to transport
Pi, ADP, and ATP across the inner membrane.

The first biochemists who studied these processes were primarily interested in the
relationship between oxygen consumption (respiration) and ATP synthesis (phosphoryla-
tion). The P/O ratio is the ratio of molecules phosphorylated to atoms of oxygen reduced.
It takes two electrons to reduce a single atom of oxygen (1/2 0 2 ) so we are interested in the
number of protons translocated for each pair of electrons that pass through complexes I,
III, and IV. Four protons are translocated by complex I, four by complex III, and two by
complex IV. Thus, for each pair of electrons that pass through these complexes from
NADH to 0 2 a total of ten protons are moved across the membrane.

Since four protons are moved back across the membrane for each molecule of cyto-
plasmic ATP, the P/O ratio isl0-^4 = 2.5. The P/O ratio for succinate is only 6 -j- 4 = 1.5
since electrons contributed by succinate oxidation do not pass through complex I.
These calculated values are close to the P/O ratios that have been observed in experiments
measuring the amount of 0 2 reduced when a given amount of ADP is phosphorylated
(Figure 14.3a). Recall that the overall energy available in the oxidation-reduction reac-
tions is 220 kj mol -1 (Section 14.4A) and this is more than enough for the synthesis of
2.5 molecules of ATP.

14.12 NADH Shuttle Mechanisms in Eukaryotes

NADH is produced by a variety of different reactions, especially the reactions catalyzed
by glyceraldehyde-3-phosphate dehydrogenase during glycolysis and those of the citric
acid cycle. NADH can be used directly in biosynthesis reactions such as amino acid
synthesis and gluconeogenesis (where glceraldehyde- 3 -phosphate dehydrogenase oper-
ates in the reverse direction).

14.12 NADH Shuttle Mechanisms in Eukaryotes

437

Excess NADH is used to produce ATP by the process that we have described in this
chapter. In bacteria, the oxidation of NADH from all sources is readily accomplished
since the membrane-associated electron transport system is embedded in the plasma
membrane and the inside surface is exposed to the cytosol. In eukaryotic cells on the
other hand, the only NADH molecules that have direct access to complex I are those
found in the mitochondrial matrix. This is not a problem for reducing equivalents pro-
duced by the citric acid cycle since that pathway is localized to the mitochondria. How-
ever, the reducing equivalents produced by glycolysis in the cytosol must enter mito-
chondria in order to fuel ATP synthesis. Because neither NADH nor NAD® can diffuse
across the inner mitochondrial membrane, reducing equivalents must enter the mito-
chondrion by shuttle mechanisms. The glycerol phosphate shuttle and malate-aspartate
shuttles are pathways by which a reduced coenzyme in the cytosol passes its reducing
power to a mitochondrial molecule that then becomes a substrate for the electron
transport chain.

The glycerol phosphate shuttle (Figure 14.26) is prominent in insect flight mus-
cles that sustain very high rates of ATP synthesis. It is also present to a lesser extent in
most mammalian cells. Two glycerol 3 -phosphate dehydrogenases are required — an
NAD® -dependent cytosolic enzyme and a membrane-embedded dehydrogenase
complex that contains an FAD prosthetic group and has a substrate binding site on
the outer face of the inner mitochondrial membrane. In the cytosol, NADH reduces
dihydroxyacetone phosphate in a reaction catalyzed by cytosolic glycerol 3 -phosphate
dehydrogenase.

CH 2 OH 3-phosphate CH 2 OH

I dehydrogenase I ^

NADH + + 0 = C < » HO — C — H + NAD^

ch 2 opo 3 © ch 2 opo 3 ©

Dihydroxyacetone Glycerol 3-phosphate (14.14)

phosphate

Glycerol 3 -phosphate is then converted back to dihydroxyacetone phosphate by the
membrane dehydrogenase complex and two electrons are transferred to the FAD prosthetic
group of the enzyme. FADH 2 transfers two electrons to the mobile electron carrier Q,
that then carries the electrons to ubiquinol: cytochrome c oxidoreductase (complex III).
The oxidation of cytosolic NADH equivalents by this pathway produces less energy
(1.5 ATP per molecule of cytosolic NADH) than the oxidation of mitochondrial NADH
because the reducing equivalents introduced by the shuttle bypass NADH:ubiquinone
oxidoreductase (complex I).

Dihydroxyacetone
NADH + H © \ , phosphate ,

Cytosolic V
glycerol 3-phosphate
dehydrogenase I

NAD^

Glycerol
3-phosphate

INTERMEMBRANE

SPACE

Glycerol 3-phosphate
dehydrogenase
complex

MATRIX

A simplified version of the malate-
aspartate shuttle is described in
Section 13.4.

◄ Figure 14.26

Glycerol phosphate shuttle. Cytosolic NADH
reduces dihydroxyacetone phosphate to glyc-
erol 3-phosphate in a reaction catalyzed by
cytosolic glycerol 3-phosphate dehydroge-
nase. The reverse reaction is catalyzed by an
integral membrane flavoprotein that trans-
fers electrons to ubiquinone.

438 CHAPTER 14 Electron Transport and ATP Synthesis

Figure 14.27 ►

Malate-aspartate shuttle. NADH in the cytosol
reduces oxaloacetate to malate that is trans-
ported into the mitochondrial matrix. The
reoxidation of malate generates NADH that
can pass electrons to the electron transport
chain. Completion of the shuttle cycle re-
quires the activities of mitochondrial and
cytosolic aspartate transaminase.

NADH,H©

NAD© I Cytosolic aspartate

J , transaminase

Malate< ^ ^ — Oxaloacetate^ —

Dicarboxylate

translocase

Cytosolic malate
dehydrogenase

a - Ketog I uta rate -

Aspartate

Glutamate

Glutamate-

aspartate

translocase

MATRIX

Glutamate

-u-Ketoglutarate^-

Mitochondrial malate

dehydrogenase , ,

Malate p— \ — » Oxaloacetate ,> Aspartate

NAD©'

NADH,H©

* Mitochondrial
aspartate
transaminase

,©

Electron transport chain
(in inner membrane)

▲ Another kind of shuttle. This one required a
great deal of energy.

The malate-aspartate shuttle is more common. This shuttle requires cytosolic versions
of malate dehydrogenase — the same enzyme used to convert cytosolic malate to ox-
aloacetate for gluconeogenesis. The reverse reaction is required for the malate-aspartate
shuttle. The operation of the shuttle is diagrammed in Figure 14.27. NADH in the cy-
tosol reduces oxaloacetate to malate in a reaction catalyzed by cytosolic malate dehydro-
genase. Malate enters the mitochondrial matrix via the dicarboxylate translocase in
electroneutral exchange for a-ketoglutarate. Inside the mitochondria, the citric acid
cycle version of malate dehydrogenase catalyzes the reoxidation of malate to oxaloac-
etate with the reduction of mitochondrial NAD® to NADH. NADH is then oxidized by
complex I of the membrane-associated electron transport chain.

Continued operation of the shuttle requires the return of oxaloacetate to the cytosol
but oxaloacetate cannot be directly transported across the inner mitochondrial membrane.
Instead, oxaloacetate reacts with glutamate in a reversible reaction catalyzed by mitochon-
drial aspartate transaminase (Section 17.7C). This reaction transfers an amino group to ox-
aloacetate producing aspartate and a-ketoglutarate. Each molecule of a-ketoglutarate
exits the mitochondrion via the dicarboxylate translocase in exchange for malate.
Aspartate exits through the glutamate-aspartate translocase in exchange for glutamate.
Once they are in the cytosol, aspartate and a-ketoglutarate become the substrates for a
cytosolic form of aspartate transaminase that catalyzes the formation of glutamate and
oxaloacetate. Glutamate re-enters the mitochondrion in antiport with aspartate and ox-
aloacetate reacts with another molecule of cytosolic NADH, repeating the cycle.

This complex shuttle system requires several enzymes that have distinctive cyto-
plasmic and mitochondrial versions (e.g., malate dehydrogenase). As a general rule,
these enzymes are encoded by different, but related, genes that are descended from a
common ancestor by an ancient gene duplication event. The compartmentalization of
metabolic pathways in eukaryotic cells provides them with some advantages over bacte-
rial cells but it requires mechanisms for moving metabolites across internal mem-
branes. Part of the cost of compartmentalization is the duplication of enzymes that
need to be present in several compartments. This partly explains why eukaryotic
genomes contain so many families of related genes while bacterial genomes usually have
only a single copy. One of the striking features of the human genome sequence is the
presence of many gene families of this sort. Another major discovery is the presence

14.13 Other Terminal Electron Acceptors and Donors 439

BOX 14.3 THE HIGH COST OF LIVING

The average active adult needs about 2400 kilocalories
(10,080 kj) per day. If all of this energy was translated to ATP
equivalents, then it would correspond to the hydrolysis of
210 moles of ATP per day. (Assuming that the Gibbs free en-
ergy of hydrolysis is 48 kj mol -1 .) This is approximately equal
to 100 kg of ATP (M r = 507).

All these ATP molecules have to be synthesized and by
far the most common pathway is the synthesis of ATP driven
by mitochondrial proton gradients. Actual calculated and
measured values suggest that the average person makes

9 X 10 20 molecules of ATP per second or 78 X 10 24 molecules
per day. This is 130 moles or 66 kg of ATP.

Thus, a significant percentage of our calorie intake is
converted into a mitochondrial proton gradient in order to
drive ATP synthesis. These calculations also tell us that ATP
molecules turn over very rapidly since our bodies don’t
contain 66 kg of ATP.

Rich, R (2003). The cost of living. Nature 421, 583.

of hundreds of genes involved in the translocation of molecules across membranes.
The dicarboxylate translocase and glutamate-aspartate translocase described here
(Figure 14.27) are examples of transport proteins.

14.13 Other Terminal Electron Acceptors and Donors

Up to this point we have only considered NADH and succinate as important sources of
electrons in membrane-associated electron transport. These reduced compounds are
mostly derived from catabolic oxidation-reduction reactions such as those in glycolysis
and the citric acid cycle. You can imagine that the ultimate source of glucose is a biosyn-
thesis pathway within a photosynthetic organism. The electrons in the chemical bonds
of glucose were put there using light energy — the energy from sunlight is ultimately
what powers ATP synthesis in mitochondria.

This is a reasonably accurate picture of energy flow in the modern biosphere but it
doesn’t explain how life survived before photosynthesis evolved. Not only did photo-
synthesis provide an abundant source of carbon compounds but it is also responsible for
the increase in oxygen levels in the atmosphere. As we will see in the next chapter, photo-
synthesis also requires a membrane-associated electron transport system coupled to ATP
synthesis. It’s quite likely that respiratory electron transport, as described in this chapter,
evolved first and the photosynthesis mechanism came later. There was probably life on this
planet for several hundred million years before photosynthesis became commonplace.

What was the ultimate source of energy before sunlight? We have a pretty good idea
of how metabolism worked in the beginning because there are still chemoautotrophic
bacteria alive today. These species do not need organic molecules as carbon or energy
sources and they do not capture energy from sunlight.

Chemoautotrophs derive their energy from oxidizing inorganic compounds such
as H 2 , NH^, NO®, H 2 S, S, or Fe©. These inorganic molecules serve as a direct source
of energetic electrons in membrane-associated electron transport. The terminal elec-
tron acceptors can be 0 2 , fumarate, or a wide variety of other molecules. As electrons
pass through their electron transport chain a protonmotive force is generated and ATP
is synthesized. An example of such a pathway is shown in Figure 14.28.

The electron donor is hydrogen in this example. A membrane-bound hydrogenase
oxidizes hydrogen to protons. Such hydrogenases are common in a wide variety of bacte-
ria species. Electrons pass through cytochrome complexes similar to those of respiratory
electron transport. In most bacteria, the mobile quinone is not ubiquinone but a related
molecule called menaquinone (Section 7.15). Fumarate reductase catalyzes the reduction
of fumarate to succinate using reduced menaquinone (MQH 2 ) as the electron donor.

E. coli can use fumarate instead of oxygen as a terminal electron acceptor when it is
growing under anaerobic conditions. Fumarate reductase is a multisubunit enzyme
embedded in the plasma membrane. It is homologous to succinate dehydrogenase and
the two enzymes catalyze a very similar reaction but in different directions. In E. coli ,

440 CHAPTER 14 Electron Transport and ATP Synthesis

▲ Figure 14.28

One possible pathway for ATP synthesis in
chemoautotrophic bacteria. Hydrogen is
oxidized by a membrane-bound hydrogenase
and electrons are passed through various
membrane cytochrome complexes. Electron
transfer is coupled to the translocation of
protons across the membrane and the re-
sulting protonmotive force is used to drive
ATP synthesis. The terminal electron accep-
tor is fumarate. Fumarate is reduced to suc-
cinate by fumarate reductase.

these two enzymes are not expressed at the same time, and in vivo each catalyzes its re-
action in only one direction (the direction related to the enzyme name). This is one of
the few cases where bacterial genomes contain a family of related genes. Each gene en-
codes a slightly different version of the same enzyme.

In addition to oxygen and fumarate, nitrate and sulfate and many other inorganic
molecules can serve as electron acceptors. There are many different combinations of
electron donors, acceptors, and electron transport complexes in chemoautotrophic bac-
teria. The important point is that these bacteria extract energy from inorganic com-
pounds in the absence of light and they may survive without oxygen.

Chemoautotrophic bacteria represent possible metabolic strategies that were pres-
ent in very ancient organisms but there are still modern bacteria that grow and repro-
duce in the absence of sunlight and oxygen such as the extreme thermophiles described
in Box 2.1 and species that live deep underground.

14.14 Superoxide Anions

One of the unfortunate consequences of oxygen metabolism is the production of reac-
tive oxygen species such as the superoxide radical (*0 2 ®), hydroxyl radical (OH*), and
hydrogen peroxide (H 2 0 2 ). All of these species are highly toxic to cells. They are pro-
duced by flavoproteins, quinones, and iron-sulfur proteins. Almost all of the electron
transport reactions produce small amounts of these reactive species, especially *0 2 ®. If
a superoxide radical is not rapidly removed by superoxide dismutase it will cause break-
down of proteins and nucleic acids.

We have already discussed superoxide dismutase as an example of an enzyme with
a diffusion controlled mechanism (Section 6.4B). The overall reaction catalyzed by this
enzyme is the dismutation of two superoxide anions to hydrogen peroxide. This reac-
tion proceeds extremely rapidly.

2 -02© + 2 H© * H 2 0 2 + 0 2 (14.15)

The rapidity of this process is typical of electron transfer reactions. In this case, a copper
ion is the only electron transfer agent bound to the enzyme. The copper ion is reduced
by superoxide anion (*0 2 ®), and it then reduces another molecule of *0 2 ®. The hy-
drogen peroxide formed can be converted to H 2 0 and 0 2 by the action of catalase.

2 H 2 0 2 > 2 H 2 0 + 0 2 (14.16)

Some bacteria species are obligate anaerobes. They die in the presence of oxygen
because they cannot deplete reactive oxygen species that arise as a by-product of oxidation-
reduction reactions. These species do not have superoxide dismutase. All aerobic species
have enzymes that scavenge reactive oxygen molecules.

Problems 441

Summary

1. The energy in reduced coenzymes is recovered as ATP through a
membrane-associated electron transport system coupled to ATP
synthesis.

2. Mitochondria are surrounded by a double membrane. The elec-
tron transport complexes and ATP synthase are embedded in the
inner membrane. This inner membrane is highly folded.

3. The chemiosmotic theory explains how the energy of a proton
gradient can be used to synthesize ATP. The free energy associated
with the protonmotive force is mostly due to the charge difference
across the membrane.

4. The electron transport complexes I through IV contain multiple
polypeptides and cofactors. The electron carriers are arranged
roughly in order of increasing reduction potential. The mobile
carriers ubiquinone (Q) and cytochrome c link the oxidation-
reduction reactions of the complexes.

5. The transfer of a pair of electrons from NADH to Q by complex I
contributes four protons to the proton concentration gradient.

6. Complex II does not directly contribute to the proton concentra-
tion gradient but rather supplies electrons from succinate oxida-
tion to the electron transport chain.

7. The transfer of a pair of electrons from QH 2 to cytochrome c by
complex III is coupled to the transport of four protons by the Q cycle.

8. The transfer of a pair electrons from cytochrome c and the reduc-
tion of 1/2 0 2 to H 2 0 by complex IV contributes two protons to
the gradient.

9. Protons move back across the membrane through complex V
(ATP synthase) . Proton flow drives ATP synthesis from ADP + Pj
by conformational changes produced by the operation of a mo-
lecular motor.

10. The transport of ADP and Pj into and ATP out of the mitochon-
drial matrix consumes the equivalent of one proton.

11. The P/O ratio, the ATP yield per pair of electrons transferred by
complexes I through IV, depends on the number of protons
translocated. The oxidation of mitochondrial NADH generates
2.5 ATP; the oxidation of succinate generates 1.5 ATP.

12. Cytosolic NADH can contribute to oxidative phosphorylation
when the reducing power is transferred to mitochondria by the
action of shuttles.

13. Superoxide dismutase converts superoxide radicals to hydrogen
peroxide. Hydrogen peroxide is removed by catalase.

Problems

1. In a typical marine bacterium the membrane potential across
the inner membrane is -0.15 V. The protonmotive force is
-21.2 kj mol -1 . If the pH in the periplasmic space is 6.35, what
is the pH in the cytoplasm if the cells are at 25°C?

2. The iron atoms of six different cytochromes in the respiratory
electron transport chain participate in one-electron transfer reac-
tions and cycle between the Fe(II) and the Fe(III) states. Explain
why the reduction potentials of the cytochromes are not identical
but range from -0.10 V to 0.39 V.

3. Functional electron transport systems can be reconstituted from
purified respiratory electron transport chain components and
membrane particles. For each of the following sets of compo-
nents, determine the final electron acceptor. Assume 0 2 is present.

(a) NADH, Q, complexes I, III, and IV

(b) NADH, Q, cytochrome c, complexes II and III

(d) succinate, Q, cytochrome c, complexes II and III

4. A gene has been identified in humans that appears to play a role
in the efficiency with which calories are utilized, and anti-obesity
drugs have been proposed to regulate the amount of the uncou-
pling protein-2 (UCP-2) produced by this gene. The UCP-2 pro-
tein is present in many human tissues and has been shown to be a
proton translocator in mitochondrial membranes. Explain how
increasing the presence of the UCP-2 protein might lead to
weight loss in humans.

5. (a) When the widely prescribed painkiller Demerol (mepiridine)

is added to a suspension of respiring mitochondria, the ratios
NADH/NAD© and Q/QH 2 increase. Which electron trans-
port complex is inhibited by Demerol?

(b) When the antibiotic myxothiazole is added to respiring mito-
chondria, the ratios cytochrome q(Fe©)/cytochrome q(Fe©)
and cytochrome fr 566 (Fe©)/cytochrome fr L ( Fe©) increase.

Where does myxothiazole inhibit the electron transport
chain?

6. (a) The toxicity of cyanide (CN©) results from its binding to the

iron atoms of the cytochrome a,a 3 complex and subsequent
inhibition of mitochondrial electron transport. How does
this cyanide-iron complex prevent oxygen from accepting
electrons from the electron transport chain?

(b) Patients who have been exposed to cyanide can be given ni-
trites that convert the Fe© iron in oxyhemoglobin to Fe©
(methemoglobin). Given the affinity of cyanide for Fe©,
suggest how this nitrite treat mentmight function to de-
crease the effects of cyanide on the electron transport chain.

7. Acyl CoA dehydrogenase catalyzes the oxidation of fatty acids.
Electrons from the oxidation reactions are transferred to FAD and
enter the electron transport chain via Q. The reduction potential
of the fatty acid in the dehydrogenase- catalyzed reaction is about
-0.05 V. Calculate the free energy changes to show why FAD — not
NAD© — is the preferred oxidizing agent.

8. For each of the following two-electron donors, state the number
of protons translocated from the mitochondrion, the number of
ATP molecules synthesized, and the P/O ratio. Assume that elec-
trons pass eventually to 0 2 , NADH is generated in the mitochon-
drion, and the electron transport and oxidative phosphorylation
systems are fully functional.

(a) NADH

(b) succinate

9. (a) Why is the outward transport of ATP favored over the outward

transport of ADP by the adenine nucleotide transporter?

(b) Does this ATP translocation have an energy cost to the
cell?

442 CHAPTER 14 Electron Transport and ATP Synthesis

10. Atractyloside is a toxic glycoside from a Mediterranean thistle
that specifically inhibits the ADP/ATP carrier. Why does atracty-
loside cause electron transport to be inhibited as well?

11. (a) Calculate the pro tonmotive force across the inner mitochon-

drial membrane at 25°C when the electrical difference is
-0.18 V (inside negative), the pH outside is 6.7, and the pH
inside is 7.5.

(b) What percentage of the energy is from the chemical (pH)
gradient, and what percentage is from the charge gradient?

12. (a) Why does NADH generated in the cytosol and transported
into the mitochondrion by the malate-aspartate shuttle pro-
duce fewer ATP molecules than NADH generated in the
mitochondrion?

(b) Calculate the number of ATP equivalents produced from the
complete oxidation of one molecule of glucose to six mole-
cules of C0 2 in the liver when the malate-aspartate shuttle is
operating. Assume aerobic conditions and fully functional
electron transport and oxidative phosphorylation systems.

Selected Readings

Mitochondria

Mentel, M., and Martin, W. (2010). Anaerobic ani-
mals from an ancient, anoxic ecological niche.
BMC Biology 8:32-38.

Taylor, R. W., and Turnbull, D. M. (2005).
Mitochondrial DNA mutations in human
disease. Nature Reviews: Genetics 6:390-402.

Chemiosmotic Theory

Lane, N. (2006) Batteries not included. Nature
441:274-277.

Mitchell, P. (1979). Keilin’s respiratory chain con-
cept and its chemiosmotic consequences. Science
206:1148-1159.

Mitchell, P., and Moyle J. (1965). Stoichiometry of
proton translocation through the respiratory
chain and adenosine triphosphatase systems of rat
liver mitochondria. Nature 208:147-151.

Schultz, B., and Chan, S. I. (2001). Structures and
proton-pumping strategies of mitochondrial res-
piratory enzymes. Annu. Rev. Biophys. Biomol.
Struct. 30:23-65.

Electron Transport Complexes

Berry, E. A., Guergova-Kuras, M., Huang, L., and
Crofts, A. R. (2000). Structure and function of
cytochrome be complexes. Annu. Rev. Biochem.
69:1005-10 75.

Brandt, U. (2006). Energy converting NADH:
quinone oxidoreductase (complex I). Annu. Rev.
Biochem. 75:69-92.

Cecchini, G. (2003). Function and structure of
Complex II of the respiratory chain. Annu. Rev.
Biochem. 72:77-100.

Clason, T., Ruiz, T., Schagger, H., Peng, G.,
Zickerman, V., Brandt, U., Michel, H., and Rader-
macher, M. (2010). The structure of eukaryotic and
prokaryotic complex I./. Struct. Biol. 169:81-88.

Clason, T., Ruiz, T., Schagger, H., Peng, G., Zicker-
man, V., Brandt, U., Michel, H., and Radermacher,
M. (2010). The structure of eukaryotic and
prokaryotic complex I. /. Struct. Biol. 169:81-88.

Crofts, A. R. (2004). The cytochrome bc x complex:
function in the context of structure. Annu. Rev.
Physiol. 66:689-733.

Hosier, J. P., Ferguson-Miller, S., and Mills, D. A.
(2006). Energy transduction: proton transfer
through the respiratory complexes. Annu. Rev.
Biochem. 75:165-187.

Hunte, C., Palsdottir, H., and Trumpower, B. L. (2003).
Protonmotive pathways and mechanisms in the
cytochrome bc\ complex. FEBS Letters 545:39-46.

Hunte, C., Zickerman, V., and Brandt, U. (2010).
Functional modules and structural basis of con-
formational coupling in mitochondrial complex I.
Science 329:448-4 57.

Richter, O.-M., and Ludwig, B. (2003). Cytochrome
c oxidase — structure, function, and physiology of
a redox-driven molecular machine. Rev. Physiol.
Biochem. Pharmacol. 147:47-74.

ATP Synthase

Capaldi, R. A., and Aggler, R. (2002). Mechanism
of the F^Q-type ATP synthase, a biological rotary
motor. Trends in Biochem. Sci. 27:154-160.

Lau, W. C. Y., and Rubinstein, J. (2010). Structure of
intact Thermus thermophilusV-ATPase by cryo-EM
reveals organization of the membrane-bound V 0

motor. Proc. Natl. Acad. Sci. (USA) 107:1367-1372.

Nishio, K., Iwamoto-Kihara, A., Yamamoto, A.,
Wada, Y., and Futai, M. (2002). Subunit rotation of
ATP synthase: a or subunit rotation relative to
the c subunit ring. Proc. Natl. Acad. Sci. (USA)
99:13448-13452.

Oster, G., and Wang, H. (2003). Rotary protein
motors. Trends in Cell Biology 13:114-121.

Other Electron Donors and Acceptors

Hederstedt, L. (1999). Respiration without 0 2 .
Science 284:1941-1942.

Iverson, T. M., Luna-Chavez, C., Cecchini, G., and
Rees, D. C. (1999). Structure of the Escherichia coli
fumarate reductase respiratory complex. Science
284:1961-1966.

Peters, J. W., Lanzilotta, W. N., Lemon, B. J., and
Seefeldt, L. C. (1998). X-ray crystal structure of the
Fe-only hydrogenase (CpI) from Clostridium pas -
teurianum to 1.8 Angstrom resolution. Science
282:1853-1858.

Tielens, A. G. M., Rotte, C., van Hellemond, J. J.,
and Martin, W. (2002). Mitochondria as we don’t
know them. Trends in Biochem. Sci. 27:564-572.

von Ballmoos, C., Cook, G. M., and Dimroth, P.
(2008). Unique rotary ATP synthase and its bio-
logical diversity .Annu. Rev. Biophys. 37:43-64.

von Ballmoos, C., Wiedenmann, A., and Dimroth,
P. (2009). Essentials for ATP synthesis by F^q ATP
synthases. Annu. Rev. Biochem. 78:649-672.

Yankovskaya, V., Horsefield, R., Tornroth, S.,
Luna-Chavez, C., Miyoshi, H., Leger, C., Byrne, B.,
and Iwata, S. (2003). Architecture of succinate de-
hydrogenase and reactive oxygen species genera-
tion. Science 299:700-704.

o c

o o
o

o
o

o o

° c

o o

Photosynthesis

T he most important part of photosynthesis is the conversion of light energy into
chemical energy in the form of ATP. The basic principle behind this fundamental
reaction is similar to that of membrane- associated electron transport covered in the
previous chapter. In photosynthesis, light shines on a pigment molecule (e.g., chlorophyll)
and an electron is excited to a higher energy level. As the electron falls back to its initial state
it gives up energy and this energy is used to translocate protons across a membrane. This
creates a proton gradient that is used to drive phosphorylation of ADP in a reaction cat-
alyzed by ATP synthase. In some cases, reducing equivalents in the form of NADPH are
synthesized directly when the excited electron is used to reduce NADP®. These reactions
are called the light reactions since they are absolutely dependent on sunlight.

Photosynthetic species use their abundant supply of cheap ATP and NADPH to
carry out all of the metabolic reactions that require energy. This includes synthesis of
proteins, nucleic acids, carbohydrates, and lipids. This is why photosynthetic bacteria
and algae are such successful organisms.

Most photosynthetic organisms have a special C0 2 fixing pathway called the Calvin
cycle. Strictly speaking, the fixation of C0 2 does not require light and is not directly
coupled to the light reactions. For this reason, these reactions are often called the dark
reactions but this does not mean they take place in the dark. This pathway is closely re-
lated to the pentose-phosphate pathway described in Section 12.4.

The details of photosynthesis reactions are extremely important in understanding
the biochemistry of all life on the planet. The ability to harvest light energy to synthesize
macromolecules led to a rapid expansion of photosynthetic organisms. This, in turn,
created opportunities for species that could secondarily exploit photosynthetic organ-
isms as food sources. Animals, such as us, ultimately derive much of their energy by de-
grading molecules that were originally synthesized using the energy from sunlight.
In addition, oxygen is a by-product of photosynthesis in plants and some bacteria.
The buildup of oxygen in Earth’s atmosphere led to its role as an electron acceptor in
membrane-associated electron transport. With few exceptions, modern eukaryotes now
absolutely depend on the supply of oxygen produced by photosynthesis in order to syn-
thesize ATP in their mitochondria.

Why does this particular group of ra-
diations , rather than some other, make
the leaves grow and the flowers burst
forth , cause the mating of fireflies
and the spawning of palolo worms ,
and, when reflecting off the surface
of the moon, excite the imagination of
poets and lovers?

Helena Curtis and Sue Barnes
(1989). Biology, 5th ed.

Top: Sunlight on trillium in the woods. Solar energy captured by photosynthetic organisms ultimately sustains the activities
of nearly all organisms on Earth.

443

444 CHAPTER 15 Photosynthesis

▲ Photosynthetic organisms. Left: cyanobac-
teria. Middle: leaves of a flowering plant.
Right: purple bacteria.

The major components of the photosynthesis reactions are large complexes of pro-
teins, pigments, and cofactors embedded in a membrane. A complex containing the
light-sensitive pigments is called a photosystem. Different species employ a variety of dif-
ferent strategies to utilize light energy in order to synthesize ATP and/or NADPH. We
will first describe the structure and function of photosystems in bacteria and then move
on to the more complex photosynthesis pathway in eukaryotes such as algae and plants.
The eukaryotic photosynthesis complexes clearly evolved from the simple bacterial ones.

15.1 Light-Gathering Pigments

There are several kinds of light- gathering pigments. They have different structures, dif-
ferent properties, and different functions.

A. The Structures of Chlorophylls

Chlorophylls are the most important pigments in photosynthesis. The structures of
the most common chlorophyll molecules are shown in Figure 15.1. Note that the
tetrapyrrole ring of chlorophylls is similar to that of heme (Figure 7.38) except that
the chlorophyll ring is reduced — it has one less double bond in the conjugated ring sys-
tem between position 7 and 8 in ring IV. Chlorophylls contain a central chelated
Mg© ion instead of the Fe© found in heme. Another distinguishing feature of chlorophylls
is that they possess a long phytol side chain that contributes to their hydrophobicity.

There are many different types of chlorophylls. They differ mostly in the side
chains labeled R l5 R 2 , and R 3 in Figure 15.1. Chlorophyll a (Chi a) and chlorophyll b

CH 3

H- CH 2 — CH — CH 2 — CH 2

— 1 3

Phytol side chain

Chi species R

Chi a

Chi b

BChl a

— CH=CH 2 — CH 3
O

-CFU — CHz,

— CH =CH 2
O

— C— CH 3

— C — H — CFU — CHz,

-CH,

— CH, — CH,

BChl b

-c— ch 3 — ch 3 — ch=ch — ch 3

Saturated in
BChl a and
BChl b

▲ Figure 15.1

Structures of chlorophyll and bacteriochlorophyll pigments. Differences in substituent groups indicated as Ri, R 2 and R 3 are shown in the table. In the
bacterioch lorophyl Is, the double bond indicated in ring II is saturated. In some molecules of bacteriochlorophyll a, the phytol side chain has three
additional double bonds. The hydrophobic phytol side chain and hydrophilic porphyrin ring give chlorophyll amphipathic characteristics. Chlorophyll
(bound to proteins) is found in photosystems and in associated light-harvesting complexes.

15.1 Light-Gathering Pigments 445

400 500 600 700

Wavelength (nm)

◄ Figure 15.2

Absorption spectra of major photosynthetic
pigments. Collectively, the pigments absorb
radiant energy across the spectrum of
visible light.

(Chi b) are found in a large number of species. Bacteriochlorophyll a (BChl a) and bac-
teriochlorophyll b (BChl b) are only found in photosynthetic bacteria. They differ from
the other chlorophylls because they have one less double bond in ring II. Pheophytin
(Ph) and bacteriopheophytin (BPh) are similar pigments where the Mg© in the central
cavity is replaced by two covalently bound hydrogens.

Chlorophyll molecules are specifically oriented in the membrane by noncovalent
binding to integral membrane proteins. The hydrophobic phytol side chain helps anchor
chlorophyll in the membrane. The light- absorbing ability of chlorophyll is due to the
tetrapyrrole ring with its network of conjugated double bonds. Chlorophylls absorb light
in the violet- to -blue region (absorption maximum 400 to 500 nm) and the orange-to-red
region (absorption maximum 650 to 700 nm) of the electromagnetic spectrum (Figure 15.2).
This is why chlorophylls are green — that’s the part of the spectrum that is reflected, not
absorbed. The exact absorption maxima of chlorophylls depend on their structures; for
example, Chi a differs from Chi b. The absorption maxima of particular chlorophyll mol-
ecules is also affected by their microenvironment within the pigment-protein complex.

KEY CONCEPT

Chlorophyll molecules are oxidized (loss
of an electron) when they absorb a
photon of light.

B. Light Energy

A single quantum of light energy is called a photon. When a chlorophyll molecule ab-
sorbs a photon, an electron from a low energy orbital in the pigment is promoted to a
higher energy molecular orbital. The energy of the absorbed photon must match the
difference in energy between the ground state and higher energy orbitals — this is why
chlorophyll absorbs only certain wavelengths of light. The excited “high energy” elec-
tron can be transferred to nearby oxidation-reduction centers in the same way that
“high energy” electrons can be transferred from NADH to FMN in complex I during
respiratory electron transport (Section 14.5). The main difference between photosyn-
thesis and respiratory electron transport is the source of excited electrons. In respiratory
electron transport the electrons are derived from chemical oxidation-reduction reactions
that produce NADH and QH 2 . In photosynthesis the electrons are directly promoted to
a “high energy” state by absorption of a photon of light.

Chlorophyll molecules can exist in three different states. In the ground state (Chi or
Chl°), all electrons are at their normal stable level. In the excited state (Chi*) a photon
of light has been absorbed. Following electron transfer, the chlorophyll molecule is
in the oxidized state (Chi®) and must be regenerated by receiving an electron from an
electron donor.

The energy of a photon of light can be calculated from the following equation

where h is Planck’s constant (6.63 x 10 34 J s), c is the velocity of light (3.00 x 10 8 m s x ),
and A is the wavelength of light. It’s often convenient to calculate the total energy of a

446 CHAPTER 15 Photosynthesis

▲ The states of chlorophyll. Reduction, exci-
tation, and oxidation of chlorophyll P680.
P680* is the excited state following absorp-
tion of a photon of light. Loss of an electron
produces the oxidized state, P680®. Gain
of an electron from an outside source (such
as the oxidation of water) yields the reduced
P680 state.

The Gibbs free energy change associ-
ated with the protonmotive force is
calculated in Section 14.3B

“mole” of photons by multiplying E by 6.022 X 10 23 (Avogadro’s number). Thus, for
light at a wavelength of 680 nm, the energy is 176 kj mol -1 . This is similar to a standard
Gibbs free energy change. It means that when a mole of chlorophyll molecules absorbs a
mole of photons the excited electrons acquire an amount of energy equal to 176 kj mol -1 .
As they fall back to their ground state they give up this energy and some of it is captured
and used to pump protons across the membrane or to synthesize NADPH.

C. The Special Pair and Antenna Chlorophylls

A typical photosystem contains dozens of chlorophyll molecules but only two special
chlorophyll molecules actually give up electrons to begin the electron transfer chain.
These two chlorophyll molecules are called the special pair. In most cases the special
pair is identified simply as pigments (P) that absorb light at a specific wavelength. Thus,
P680 is the special pair of chlorophyll molecules that absorbs light at 680 nm (red). Its
three states are P680, P680*, and P680®. P680 is the ground state. P680* is the state fol-
lowing absorption of a photon of light when the chlorophyll macromolecules have an
excited electron. P680© is the electron-deficient (oxidized) state following transfer of
an electron to another molecule. P680© is reduced to P680 by transfer of an electron
from an electron donor.

In addition to the special pair there are other specialized chlorophyll molecules that
function as part of the electron transfer chain. They accept electrons from the special
pair and transfer them to the next molecule on the pathway. Not all chlorophylls are di-
rectly involved in electron transfer. The remaining chlorophylls act as antenna mole-
cules by capturing light energy and transferring it to the special pair. These antenna
chlorophylls are much more numerous than the molecules in the electron transfer
chain. The mode of excitation energy transfer between antenna chlorophylls is called
resonance energy transfer. It does not involve the movement of electrons. You can think
of excitation energy transfer as a transfer of vibrational energy between adjacent
chlorophyll molecules in the densely packed antenna complex.

Figure 15.3 illustrates the transfer of excitation energy from antenna chlorophylls
to the special pair in one of the photosystems. The figure shows only a few of the many
antenna molecules surrounding the special pair. All chlorophyll molecules are held in

Figure 15.3 ►

Transfer of light energy from antenna chlorophyll
pigments to the special pair of chlorophyll
molecules. Light can be captured by the an-
tenna pigments (gray) and excitation energy
is transferred between antenna chlorophylls
until it reaches the special pair of chlorophyll
molecules in the electron transfer pathway
(green). The path of excitation energy trans-
fer is shown in red. The special pair gives up
an electron to the electron transfer pathway.
The chlorophyll molecules are held in fixed
positions because they are tightly bound to
membrane proteins (not shown).

15.1 Light-Gathering Pigments 447

BOX 15.1 MENDEL’S SEED COLOR MUTANT

One of Gregor Mendel’s original mutants affected the color
of the peas in a pod. The normal color of mature seeds is yel-
low (I) and the recessive mutant confers a green color to the
seeds (i). The mutation affects the “stay-green” ( sgr ) gene that
encodes a chloroplast protein responsible for the degradation
of chlorophyll as the seeds mature. When the protein is de-
fective, chlorophyll is not broken down in the chloroplasts
and the seeds stay green.

In normal wild-type plants (II) the seed are yellow and
in the heterozygotes (Ii) the deficiency in the amount of
chlorophyll degradation protein is not sufficient to affect
chlorophyll breakdown. The seeds of the heterozygotes are
also yellow. In homozygous mutant plants (ii) chlorophyll is
not degraded and the seeds are green. Mendel determined
that the wild-type trait (I) was dominant and the mutant
trait (i) was recessive. Crosses between heterozygotes (Ii x Ii)
gave the famous 3:1 ratio of yellow seeds to green seeds.

Some strains of food plants are homozygous for muta-
tions in the genes that break down chlorophyll. These “cos-
metic stay- greens,” such as the one used by Mendel, produce
seeds and fruit that are more attractive to consumers.

All the peas that we buy in supermarkets and farmer’s
markets have been genetically modified (by breeding) to be
homozygous for the deficient sgr allele. That’s why we never
see the “normal” yellow peas.

► Normal mature peas turn yellow in color as
they mature (bottom) but a mutation causes
the seeds to retain their green color (top).
The seed coat has been removed from the
lower pair of each group in order to make
the color difference more obvious.

fixed positions through interactions with the side chains of amino acids in the polypep-
tides of the photosystem. Excitation energy is efficiently transferred from any molecule
that absorbs a photon because these molecules are so close to each other.

D. Accessory Pigments

Photosynthetic membranes contain several accessory pigments in addition to chloro-
phyll. The carotenoids include /J-carotene (Figure 15.4) and related pigments such as
xanthophylls. Xanthophylls have extra hydroxyl groups on the two rings. Note that the
carotenoids, like chlorophyll, contain a series of conjugated double bonds allowing
them to absorb light. Their absorption maxima lie in the blue region of the spectrum,
which is why carotenoids appear red, yellow, or brown (Figure 15.2). The autumn colors
of deciduous trees are due, in part, to carotenoids, as is the brown color of sea kelp
(brown algae).

°ooc coo 0

▲ The autumn colors of the leaves are due,
in part, to the presence of accessory
carotenoid pigments that become visible
when chlorophyll molecules are degraded as
the leaves die.

◄ Figure 15.4

Structures of some accessory pigments.

/1-Carotene is a carotenoid, and phycoery-
thrin and phycocyanin are phycobilins. Phy-
cobilins are covalently attached to proteins
whereas carotenoids are bound noncovalently.

448 CHAPTER 15 Photosynthesis

▲ Red tide. This red tide off the coast of Fujian,
China, is due to the presence of red algae.

▲ Scytonema— a blue-green cyanobacterium.

The structure of the photosystem of the
purple bacterium, Rhodopseudomas
viridis , is shown in Figure 4.25f.

Carotenoids are closely associated with chlorophyll molecules in antenna com-
plexes. They absorb light and transfer excitation energy to adjacent chlorophylls. In ad-
dition to serving as light- gathering pigments carotenoids also play a protective role in
photosynthesis. They take up any electrons that are accidently released from antenna
chlorophylls and return them to the oxidized chlorophyll molecule. This quenching
process prevents the formation of reactive oxygen species such as the superoxide radical
(•02^). If allowed to form, these reactive oxygen species can be highly toxic to cells as
described in Section 14.14.

Phycobilins, such as red phycoerythrin and blue phycocyanin (Figure 15.4), are
found in some algae and cyanobacteria. They resemble a linear version of chlorophyll
without the central magnesium ion. Like chlorophylls and carotenoids, these molecules
contain a series of conjugated double bonds that allow them to absorb light. Like
carotenoids, the absorption maxima of phycobilins complement those of chlorophylls
and thus broaden the range of light energy that can be absorbed. In most cases, the phy-
cobilins are found in special antenna complexes called phycobilisomes. Unlike other pig-
ment molecules, the phycobilins are covalently attached to their supporting polypep-
tides. The bluish color of blue-green cyanobacteria and the red color of red algae are due
to the presence of numerous phycobilisomes associated with their photosystems.

15.2 Bacterial Photosystems

We begin our discussion by describing simple bacterial systems. These simple systems
evolved into more complicated structures in the cyanobacteria. The cyanobacterial ver-
sion of photosynthesis was then adopted by algae and plants when a primitive
cyanobacterium gave rise to chloroplasts.

Photosynthetic bacteria contain typical light-gathering photosystems. There are
two basic types of photosystems that appear to have diverged from a common ancestor
more than two billion years ago. Both types of photosystem contain a large number of
antenna pigments surrounding a small reaction center located in the middle of the
structure. The reaction center consists of a few chlorophyll molecules that include the
special pair and others forming a short electron transfer chain.

Photosystem I (PSD contains a type I reaction center. Photosystem II (PSII) contains a
type II reaction center. Heliobacteria and green sulfur bacteria rely on photosystems with
a type I reaction center whereas purple bacteria and green filamentous bacteria use
photosystems with a type II reaction center. Cyanobacteria, the most abundant class of
photosynthetic bacteria, utilize both photosystem I and photosystem II coupled in se-
ries. This coupled system resembles the one found in algae and plants.

A. Photosystem II

We begin by describing photosynthesis in purple bacteria and green filamentous bacteria.
Most of these species of bacteria are strict anaerobes — they cannot survive in the presence
of oxygen. Thus, they do not produce oxygen as a by-product of photosynthesis or con-
sume it in respiratory electron transport. Purple bacteria and green filamentous bacteria
have photosystems with a type II reaction center. These membrane complexes are often
referred to as the bacterial reaction center (BRC) but this is misleading since bacteria
also contain the other type of reaction center. We will refer to it here as photosystem II
since it is evolutionarily related to photosystem II in cyanobacteria and eukaryotes.

The structure of the purple bacteria photosystem is shown in Figure 15.5. The pig-
ment molecules of the internal type II reaction center form an electron transfer chain
with two branches. The special pair of bacteriochlorophylls (P870) are positioned near
the periplasmic (outside) surface of the membrane. Each branch contains a molecule of
bacteriochlorophyll a and a bacteriopheophytin molecule (Figure 15.6). The right-hand
branch terminates in a tightly bound quinone molecule while the equivalent position in
the left-hand branch is occupied by a loosely bound quinone that can dissociate and
diffuse within the lipid bilayer. Note in Figure 15.5 that the bound quinone is buried
within the a helix barrel spanning the membrane while the equivalent site on the other
side of the complex is open to the lipid bilayer.

15.2 Bacterial Photosystems 449

OUTSIDE

(Periplasm)

Bacterial

membrane

INSIDE

(Cytoplasm)

Cytochrome c

Electron transfer begins with the release of an excited electron from P870 following
absorption of a photon of light or the transfer of excitation energy from antenna pig-
ments. (Antenna pigment molecules are not shown in Figure 15.6.) Electrons are then
transferred exclusively down the right-hand branch of the reaction center complex
resulting in the reduction of the bound quinone molecule. From there, electrons are
passed to the mobile quinone on the opposite side of the complex. This transfer is
mediated by a single bound iron atom on the central axis near the cytoplasmic side of
the membrane. The mobile quinone (Q) is reduced to QH 2 in a two-step process via the
sequential transfer of two electrons and the uptake of two H© from the cytoplasm. Two
photons of light are absorbed for each molecule of QH 2 produced. Modern type II reac-
tion centers probably evolved from a more primitive system in which electrons were
transferred down both branches to produce QH 2 at both of the Q sites.

QH 2 diffuses within the lipid bilayer to the cytochrome bc\ complex (complex III)
of the bacterial respiratory electron transport system. This is the same complex that we
described in the previous chapter (Section 14.7). The cytochrome bc\ complex catalyzes
the oxidation of QH 2 and the reduction of cytochrome c — the enzyme is ubiquinol:
cytochrome c oxidoreductase. This reaction is coupled to the transfer of H© from the
cytoplasm to the periplasmic space via the Q cycle. The resulting proton gradient drives
the synthesis of ATP by ATP synthase (Figure 15.7).

The P870© special pair of chlorophyll molecules is reduced by the cytochrome c
(Fe@) molecules produced by the cytochrome bc\ complex. Cytochrome c diffuses
within the periplasmic space enclosed by the two membranes surrounding the bacterial
cell. The net effect is that electrons are shuffled from PSII to the cytochrome bci com-
plex and back again. Note that the structure shown in Figure 15.5 includes a bound cy-
tochrome c molecule with its heme group positioned near the P870 special pair in order
to facilitate electron transfer.

The movement of electrons between complexes is mediated by the mobile cofactors
QH 2 and cytochrome c just as we saw in respiratory electron transport. The main differ-
ence between photosynthesis in purple bacteria and respiratory electron transport is
that photosynthesis is a cyclic process. There is no net gain or loss of electrons to other
reactions and consequently no outside source of electrons is needed. Cyclic electron
flow is a characteristic of many, but not all, photosynthesis reactions. The result of cou-
pling PSII and the cytochrome bc\ complex is that absorption of light creates a proton

◄ Figure 15.5

Photosystem II in the purple bacterium
Rhodobacter spaeroides. The core of the
structure consists of two homologous
membrane-spanning polypeptide subunits
(L and M). Each subunit has five transmem-
brane a helices. The electron transfer mole-
cules of the reaction center are sandwiched
between the core polypeptides. Cytochrome
c binds to PSII on the periplasmic side of
the membrane (top). An additional subunit
covers the core subunits on the cytoplasmic
surface (bottom). [PDB 1L9B]

Bound

quinone

Special pair
(P870)
Bacteria
chloro

Cytochrome
c heme

hv (x2)

▲ Figure 15.6

The type II reaction center contains the elec-
tron transfer chain. The special pair (P870)
is located near the periplasmic surface close
to the heme group of cytochrome c. When
light is absorbed, electrons are transferred
one at a time from P870 to BChl a to BPh
to a bound quinone and from there to a
quinone located at a loosely bound site next
to a central iron atom (orange). Electrons
are restored to P870 from cytochrome c.

450 CHAPTER 15 Photosynthesis

Figure 15.7 ►

Photosynthesis in purple bacteria. Light is
absorbed by the pigments of the PSII com-
plex resulting in the transfer of electrons
from P870 to QH 2 via the reaction center
electron transfer chain. QH 2 diffuses to the
cytochrome bc\ complex where the elec-
trons are transferred to cytochrome c. This
reaction is coupled to the transfer of protons
across the membrane. The proton gradient
drives the synthesis of ATP. Reduced cy-
tochrome c diffuses within the periplasmic
space to PSII where it reduces P870 + . The
Q-cycle reactions are shown in more detail
in Figure 14.11.

PSII

Cytochrome be ^
complex

ADP + Pj ATP

KEY CONCEPT

Bacteria with photosystem II use sunlight
to produce a proton gradient that drives
ATP synthesis.

KEY CONCEPT

Photosynthesis in purple bacteria is a
cyclic process. It does not require an
external source of electrons such as
H 2 0 or H e S.

gradient for ATP synthesis. The reactions are listed in Table 15.1. (The cytochrome bc x
reactions are the same ones shown in Table 14.3.) Four protons are transferred across
the membrane for every two photons of light that are absorbed. The ATP molecules
produced as a result of this cycle are used by bacteria to synthesize proteins, nucleic
acids, carbohydrates, and lipids. Thus, captured light energy is ultimately used in
biosynthesis reactions.

We can calculate the energy of two “moles” of light at 870 nm using Equation 15.1.
It works out to 274 kj mol -1 . This light energy is used to pump four protons across the
membrane. Pumping requires approximately 4 X 19.4 kj mol -1 = 77.6 kj mol -1 using
our estimate from the previous chapter (Section 14.3). The result suggests that the pro-
duction of chemical energy from light energy is not very efficient in purple bacteria
(77.6/274 = 28%).

The basic principle of photosynthesis is the conversion of light energy (photons) to
chemical energy (e.g. ATP). The pathway clearly evolved, in part, from the electron
transport system we described in the previous chapter. Photosynthesis evolved several
hundred million years after the main energy-producing pathway that uses complex III
and ATP synthase. It’s important to note that the ATP produced in bacterial photosyn-
thesis is not restricted to the synthesis of carbohydrate and oxygen is not produced as
part of the process.

Table 15.1 Photosystem II reactions

PSII: 2 P870 + 2 photons > 2 P870© + 2 e 0

Q + 2 e 0 + 2 H 0 in > QH 2

Cyt be y. 2 QH 2 + 2 cyt c (Fe©) > 2 Q + 2 cyt c (Fe©) + 4 H 0 out

Q + 2 e 0 + 2 H 0 in > QH 2

PSII: 2 cyt c (Fe© ) + 2 P87O 0 * 2 cyt c (Fe© ) + 2 P870

Sum: 2 photons + 4 H 0 in > 4 H 0 out

+ 2 e 0

B. Photosystem I

The structure of a typical photosystem I (PSI) complex is shown in Figure 15.8. The
central part of the complex is formed by two homologous polypeptides with multiple
membrane-spanning a helices. Each subunit of this dimer has two domains — an inte-
rior domain that binds the electron transfer chain pigments of the type I reaction center
and a peripheral domain that binds antenna pigments. The reaction center protein do-
mains in PSI subunits are related by structure and amino acid sequence to the core
polypeptides in PSII. This is strong evidence for a common ancestor of type I and type
II reaction centers.

15.2 Bacterial Photosystems 451

OUTSIDE

(Periplasm)

Bacterial

membrane

INSIDE

(Cytoplasm)

The most obvious difference between PSI and PSII is the presence of a more complex
antenna structure in PSI than in PSII. The PSI antenna complex is packed with chloro-
phyll and carotenoid pigment molecules. The example shown in Figure 15.8 is from
cyanobacteria whose PSI complexes contain 96 chlorophylls and 22 carotenoids. Many of
the light-gathering pigment molecules are tightly bound to additional membrane-
spanning polypeptide subunits that surround the core subunits. The contrast between
the structures shown in Figure 15.5 and Figure 15.8 is a bit misleading since there are sim-
pler forms of PSI in some bacteria and more complex versions of PSII in other species (see
below). Nevertheless, as a general rule, PSI is larger and more complicated than PSII.

The organization of the electron transfer chain molecules in PSI reveals striking par-
allels to that of PSII (Figure 15.9). In both cases, the reaction center contains two short
branches of pigment molecules that terminate at bound quinones. The PSI pigment mol-
ecules are both chlorophylls and not one chlorophyll and one pheophytin as in PSII. The
bound quinones in PSI are usually phylloquinones whereas in PSII they are related to
ubiquinone (or menaquinone in bacteria). The phylloquinones in type I reaction centers
are tightly bound to the complex and form part of the electron transfer chain. (Recall that
one of the quinones in type II reaction centers is a mobile terminal electron acceptor.)

Electron transfer begins with a special pair of chlorophyll molecules located near
the periplasmic surface of the membrane. This special pair is known as P700 since it ab-
sorbs light at a wavelength of 700 nm. The two chlorophyll molecules are not identical —
the molecule closest to the A-branch is an epimer of chlorophyll a (bacteriochlorophyll a
in bacteria). P700 is excited by absorbing a photon of light or by excitation energy
transfer from antenna molecules. The excited electron is then transferred down one of
the branches of the electron transfer chain to one of the bound phylloquinones. Electron
transfer from P700 to phylloquinone takes about 20 picoseconds (1(T 12 s). This is extremely
rapid compared to other electron transfer systems. In type II reaction centers, for example,
the transfer from P680 to the bound quinone takes two or three times longer.

Electrons are subsequently transferred from bound phylloquinone to the three Fe-S
clusters, F x , F A , and F B . The terminal electron acceptor in PSI is ferredoxin (or flavodoxin)
(Figure 7.36). Ferredoxin contains two [4Fe-4S] iron-sulfur clusters and reduction in-
volves a Fe© —> Fe© reduction with a standard reduction potential of —0.43 V (Table 10.5).

Reduced ferredoxin (Fd re( j) becomes the substrate for an oxidation-reduction reac-
tion catalyzed by an enzyme called ferredoxin:NADP® oxidoreductase, more com-
monly known as ferredoxin:NADP® reductase or FNR. The enzyme is a flavoprotein
(containing FAD) and the reaction proceeds in three steps involving a typical semi-
quinone intermediate (Section 7.5). The product of the reaction is reducing equivalents
in the form of NADPH. The coupled reactions involving PSI are shown in Table 15.2.

Note that the standard reduction potential of ferredoxin is considerably lower than
that ofNADP®, allowing for transfer of electrons from ferredoxin to NDAP© . The
terminal electron acceptor is Q in photosystem II and its standard reduction potential is

◄ Figure 15.8

Structure of photosystem I (PSI). This version
of PSI is from the cyanobacterium Therm-
osynechococcus elongatus ( Synechococcus
elongatus ). The complex contains 96 chloro-
phylls (green), 22 carotenoids (red), and
three iron-sulfur clusters (orange). There
are 14 polypeptide subunits, most of
which have membrane-spanning a helices.
[PBD 1JB0]

Phylloquinone is also known as vitamin K
(Section 7.14D, Figure 7.29).

e ©

A-branch

Ferredoxin

Flavodoxin

P700

Cytochrome c
or

Plastocyanin

▲ Figure 15.9

PSI electron transfer chain (type I reaction
center). Electron transfer begins with the
special pair of chlorophyll molecules (P700)
and proceeds down one of the branches to
phylloquinone. From there, electrons are
transferred to the Fe-S clusters and eventu-
ally to ferredoxin. P700© is reduced by
cytochrome cor plastocyanin.

452

CHAPTER 15 Photosynthesis

KEY CONCEPT

Bacteria with photosystem I use sunlight
to produce NADPH.

Ferredoxin (Fe@) + e® — > Fe©

A£ = -0.43 V

NADP® + H® + 2 e 0 -> NADPH
A E= -0.32 V

Ubiquinone (Q) + 2 H® +2 e 0 — » QH 2
A E = +0.04 V

▲ Green sulfur bacteria. Agar plate with
streaks of Chlorobium tepidum.

Table 15.2 The photosystem I reactions

PSI:

FNR:

2 P700 + 2 photons > 2 P700© + 2 e©

2 Fd ox + 2 e© + >2 Fd red

Fd red + H© + FAD Fd ox + FADH-

Fd red + H© + FADH- Fd ox + FADH 2

FADH 2 + NADP© FAD + NADPH + H©

Sum: 2 P700 + 2 photons + NADP© + H© > 2 P700© + NADPH

too high to allow transfer of electrons to NADP®. This means that energy capture from
sunlight is more efficient in PSII than in PSI.

The reactions in PSI do not create a cyclic pathway. The oxidized special pair in
type I reaction centers (P700® ) must be reduced by electrons from an outside source since
the excited chlorophyll electrons were eventually transferred to NADPH. Some bacteria
contain versions of PSI that bind cytochrome c on the outside surface of the membrane
next to the special pair. In these bacteria P700® is reduced by reduced cytochrome c in
a manner similar to the reduction of the special pair in PSII. The source of electrons for
reduced cytochrome c depends on the species. In green sulfur bacteria it is various
reduced sulfur compounds such as H 2 S and S 2 0©. The oxidation of these sulfur com-
pounds is coupled to the transfer of electrons to cytochrome c by special enzymes that
are found in these species (Figure 15.10). Green sulfur bacteria are photoautotrophs
(Section 10.3) that grow in the absence of oxygen.

Noncyclic electron transfer is a characteristic feature of PSI but there can also be a
cyclic process of electron transfer. Some electrons from PSI are occasionally passed from
ferredoxin to a quinone — probably by ferredoximquinone oxidoreductase (ferredoxin:
quinone reductase, FQR). Quinol (QH 2 ) interacts with the cytochrome frq complex

▲ Figure 15.10

Photosynthesis in green sulfur bacteria. Photoactivation of P700 leads to production of reduced
ferredoxin on the cytoplasmic side of the membrane. Ferredoxin becomes the electron donor in a
reaction catalyzed by ferredoximNADP© reductase (FNR) resulting in production of NADPH in the
cytoplasm. Ferredoxin can also reduce Q to QH 2 in a reaction catalyzed by ferredoximquinone
reductase (FQR). QH 2 is oxidized by the cytochrome bc\ complex, resulting in the transfer of elec-
trons to reduced cytochrome c and the transfer of protons across the membrane. P700© is nor-
mally reduced by cytochrome c on the periplasmic side of the membrane. In the noncyclic process,
reduced cytochrome c is made in reactions that are coupled to the oxidation of sulfur compounds
such as H 2 S. The transfer of electrons is shown by red arrows.

15.2 Bacterial Photosystems 453

transferring electrons via cytochrome bc x to cytochrome c and cytochrome c reduces
P700© (Figure 15.10). This cyclic process is very similar to the coupled reactions involv-
ing PSII. It allows for light-mediated synthesis of ATP because the passage of electrons
through cytochrome bc x is associated with the translocation of protons across the mem-
brane via the Q cycle. In most cases, the noncyclic process predominates and NADPH is
produced; however, if NADPH cannot be efficiently used in biosynthesis reactions, elec-
trons will be transferred through cytochrome bc x to produce ATP.

C. Coupled Photosystems and Cytochrome bf

Cyanobacterial membranes contain both PSI and PSII. The two photosystems are coupled
in series to produce both NADPH and ATP in response to light. The photosynthetic reac-
tions in cyanobacteria are illustrated in Figure 15.1 1. Light is absorbed by PSII leading to
excitation of P680 and transfer of an electron to a mobile quinone called plastoquinone
(PQ, Figure 7.33). Electrons are then transferred to a cytochrome £/ complex similar to
the cytochrome bc x complex in respiratory electron transport. Electron transport within
the cytochrome bf complex is coupled to the movement of H© across the membrane by a
photosynthetic Q cycle. The coupling of PSII and a cytochrome £/ complex is similar in
principle to photosynthesis reactions in purple bacteria with one major difference — in
purple bacteria electrons are returned to PSII by the terminal electron acceptor of the
cytochrome bc x complex (cytochrome c) whereas in cyanobacteria electrons are passed on
to PSI. The terminal electron acceptor of the unique cytochrome bf complex is either
cytochrome c or a blue copper-containing protein called plastocyanin (PC). Reduced cy-
tochrome c and reduced plastocyanin are mobile carriers that bind to the outside (periplas-
mic) surface of PSI and reduce P700©. (Most cyanobacteria and algae use cytochrome c
while some cyanobacteria and all plants use plastocyanin, or a different cytochrome called
cytochrome c 6 , as the terminal electron acceptor of the cytochrome //complex.)

The structure of the photosynthetic cytochrome bf complex has been solved by
X-ray crystallography (Figure 15.12). It contains a cytochrome b with two cytochrome
reaction centers whose role in the Q cycle is similar to that of cytochrome b in the
cytochrome Z?q complex (complex III) of respiratory electron transport. A Rieske
iron-sulfur protein (ISP) transports electrons from one of the cytochrome b sites to cy-
tochrome/and reduced cytochrome /passes electrons to plastocyanin. Cytochrome/
(/stands for feuille, the French word for leaf) is a distinct protein unrelated to cytochrome
c 1 of the respiratory cytochrome bc 1 complex but cytochrome b and ISP are homo-
logues of the proteins found in complex III.

The cytochrome bf complex evolved from the original cytochrome bc\ complex
that was present in ancient cyanobacteria. The most important adaptation was the
replacement of cytochrome c 1 of the bacterial be complex with cytochrome /in the
cyanobacterial complex. This change allowed for the transfer of electrons to the copper-
containing plastocyanin via cytochrome/. (Recall that mobile cytochrome c,
not plastocyanin, is the normal electron acceptor of the cytochrome bc x complex.)

Reduced ferredoxin can be used directly
in other pathways, notably in nitrogen
fixation (Section 17.1)

KEY CONCEPT

Organisms with coupled photosystem I
and photosystem II use sunlight to
produce both NADPH a/icf a proton
gradient that drives ATP synthesis.

v Figure 15.1 1

Photosynthesis in cyanobacteria. Light (wavy
arrows) is captured and used to drive the
transport of electrons (obtained from water)
from PSII through the cytochrome bf com-
plex to PSI and ferredoxin. This process can
generate NADPH and a proton concentration
gradient that is used to drive phosphoryla-
tion of ADP. For each water molecule oxi-
dized to 1/2 0 2 by the oxygen evolving com-
plex (OEC), one molecule of NADP© is
reduced to NADPH. For simplicity, PSI,

PSII, and cytochrome bf are shown close
together in the plasma membrane but in
most species they are located within internal
membrane structures. Plastoquinone (PQ) is
the mobile carrier between PSII and the cy-
tochrome bf complex. In this example, plas-
tocyanin (PC) is the mobile carrier between
the cytochrome bf complex and PSI.

INSIDE

Cytochrome bf cyclic '' 2 Fd ri /NADP+ + H 4
complex electron j{ FNR

transfer

2 Fd n

^ NADPH ADP + P; ATP

454 CHAPTER 15 Photosynthesis

Figure 15.12 ►

Cytochrome complex from the cyanobac-
terium Mastigocladus laminosus. The complex
contains two functional enzymes as in com-
plex III (compare Figure 14.10). The pri-
mary electron transfer components are:
heme b L and heme b H (the sites of Q-cycle
oxidation reactions), the iron-sulfur cluster
(Fe-S) in ISP, and heme f. Each unit also
contains a chlorophyll a, a /1-carotene, and
an unusual heme x whose function is
unknown (not shown). [PDB 1UM3]

Heme f
Fe-S

Heme b L
Heme b H

KEY CONCEPT

The splitting of water to form molecular
oxygen arose in order to supply electrons
to photosystem II.

Plastocyanin binds specifically to PSI in cyanobacteria and transfers electrons to P700® .
This allows for a unidirectional flow of electrons from PSII — > PQH 2 —> cytochrome
&/-> PC PSI -> NADPH.

Cyanobacteria do not contain cytochrome 2?q. Thus, cytochrome bf also plays a
role in respiratory electron transport because it replaces the normal complex III. Re-
duced plastocyanin is the electron donor to the terminal oxidase (complex IV) possibly
via an intermediate cytochrome c - like carrier. Plastoquinone is the mobile quinone
electron carrier in both photosynthesis and respiratory electron transport.

Photoactivation of PSI results in synthesis of NADPH in a manner similar to that
in green sulfur bacteria. As in green sulfur bacteria, some electrons are recycled but in
this case it is through the cytochrome bf complex. Note that PSII, cytochrome bf and
PSI are coupled in series and the transfer of electrons to NADPH results in a deficiency
of electrons at P680® in PSII. The reduction of P680® in cyanobacteria is accom-
plished by extracting electrons from water with the production of oxygen as a by-
product. The enzyme that splits water is called the oxygen evolving complex (OEC) and
it is tightly bound to PSII on the outer surface of the membrane. The evolution of an
oxygen evolving complex in primitive cyanobacteria was one of the most important
biochemical events in the history of life.

The oxygen evolving complex (OEC) contains a cluster of Mn® ions, a Ca® ion,
and a Cl® ion. It catalyzes a complex reaction in which four electrons are extracted, one
at a time, from two molecules of water. The reaction takes place on the outside of the
PSII complex near the special pair of chlorophyll molecules (P680). The electrons from
the splitting of water are transferred to P680® (Figure 15.13). The exact mechanism of
the water splitting reaction is being investigated in a number of laboratories. It is simi-
lar, in principle, to the reverse reaction catalyzed by complex IV of the respiratory elec-
tron transport chain (Section 14.8). Note that the oxygen evolving complex is located
on the exterior surface of the membrane and the release of protons from water con-
tributes to the formation of the proton gradient across the membrane.

As mentioned earlier, the similarities between PSI and PSII indicate that they
evolved from a common ancestor. Over time, these two photosystems diverged in those
species of photosynthetic bacteria that contain only one of the two types (e.g., purple
bacteria, green sulfur bacteria). At some point, about 2.5 billion years ago, a primitive
ancestor of cyanobacteria acquired both types of photosystem — probably by taking up
a large part of the genome from an unrelated bacterial species. At first the two types of

15.2 Bacterial Photosystems 455

Oxygen Evolving
Complex

◄ Figure 15.13

PSII and the oxygen-evolving center. The PSII

complex in the cyanobacterium Thermosyne-
chococcus elongatus is much larger than the
PSII complex in purple bacteria (Figure 15.5)
but the core structures are very similar. The
cyanobacteria complex contains many an-
tenna chorolophylls and carotenoids and
it is a dimer. The oxygen evolving complex
(OEC) contains a Mn 3 Ca0 4 cluster (circled)
where the splitting of water occurs. This
metal ion cluster is positioned over the type
II reaction center. [PDB 3BZ1]

photosystem must have worked in parallel but they began to function in series with the
evolution of a photosynthetic cytochrome Z?/ complex (from cytochrome bc{) and an
oxygen evolving complex. Later on, a species of cyanobacteria entered into a symbiotic
relationship with a primitive eukaryotic cell and this led to the modern chloroplasts
found in algae and plants.

The coupled photosystems are able to capture light energy and use it to produce
both ATP (from the proton gradient) and reducing equivalents in the form of NADPH.
Neither photosystem by itself can accomplish these two goals with the same efficiency.

The net result of this simplified linear pathway is the production of one molecule
of NADPH and the transfer of four protons across the membrane for each pair of elec-
trons excited by the absorption of light energy in each photosystem. The two separate
excitation steps in PSI and PSII require a total of four photons of light energy. The split-
ting of water by the OEC contributes to the proton gradient and produces molecular
oxygen. The individual reactions are summarized in Table 15.3.

D. Reduction Potentials and Gibbs Free Energy in Photosynthesis

The path of electron flow during photosynthesis can be depicted in a zigzag figure
called the Z-scheme (Figure 15.14). The Z-scheme plots the reduction potentials of the
photosynthetic electron transfer components in PSI, PSII, and cytochrome bf It shows
that the absorption of light energy converts P680 and P700 — pigment molecules that
are poor reducing agents — to excited molecules (P680* and P700*) that are good

Table 15.3 The photosynthesis reactions in species with both photosystems

PSII:
OEC:
Cyt bf:
PSI:

FNR:

2 P680 + 2 photons » 2 P680© + 2 e Q

PQ + 2 e 0 + 2 H© in > PQH 2

H 2 0 * \o 2 + 2 H© out + 2 e©

2 P680© + 2 e© > 2 P680

2 PQH 2 + 2 plastocyanin (Cu©) * 2 PQ + 2 plastocyanin (Cu©) + 4 H© out + 2 e©

PQ + 2 H© in + 2c© PQH 2

2 P700 + 2 photons » 2 P700© + 2 e©

2 Fd ox + 2 e© >2 Fd red

2 plastocyanin (Cu©) + 2 P700© * 2 plastocyanin (Cu 2+ ) + 2 P700

2 Fd re d + H© + NADP© 2 Fd ox + NADPH

Sum: H 2 0 + 4 photons + 4 H© in + NADP© + H© * ^0 2 + 6 H© out + NADPH

456

CHAPTER 15 Photosynthesis

KEY CONCEPT

The energy from a photon of light is used
to excite an electron in the special pair of
chlorophyll molecules. The excited state
has a much lower reduction potential
making it easy to give up an electron to
an oxidation reaction.

reducing agents. (Recall that a reducing agent is one that gives up electrons to reduce
another molecule. The reducing agent is oxidized in such reactions.) The oxidized
forms of the pigment molecules are P680© and P700©. Energy is recovered when
P680* and P700* are oxidized and electrons are passed to cytochrome bf and NADPH.

The standard reduction potentials of many of these components are listed in
Table 10.5. The difference between any two reduction potentials can be converted to a
standard Gibbs free energy change as we saw in Chapter 10. Looking at Figure 15.14 we
can see that the absorption of a photon by either P680 or P 700 lowers the standard re-
duction potential by about 1.85 V. In these examples, a difference of 1.85 V corresponds
to a standard Gibbs free energy change of about 180 kj mol -1 (AG°' = 180 kj mol -1 ).
This value is almost identical to the calculated energy of a “mole” of photons at a wave-
length of 680 nm (176 kj mol -1 , Section 15.1). What this means is that the energy of
sunlight is very efficiently converted to a change in reduction potential.

There are many similarities between electron transfer in photosynthesis and the
membrane-associated electron transport chain that we saw in the last chapter. In both cases
electrons pass through a cytochrome complex that transports H© across a membrane.
The resulting proton gradient is expended when ATP is synthesized by ATP synthase.

The structure and orientation of cytochrome bc x (complex III) and cytochrome bf
are similar. Both complexes release protons into the space between the inner and outer
membranes. The orientation of ATP synthase is also identical — the “head” of the struc-
ture is located in the cytoplasm of bacterial cells or the inside compartment of mito-
chondria. In the next section we’ll see that the orientation of ATP synthase in chloroplasts
is topologically similar.

-1.5 -|

PSI

- 1.0 -

-0.5 -

0 -

+0.5 -

+ 1.0 -

+1.5 - 1

P700*

▲ Figure 15.14

Z-scheme, showing reduction potentials and electron flow during photosynthesis in cyanobacteria. Light energy is absorbed by the special pair pigments,
P680 and P700. This converts these molecules into strong reducing agents as shown by the huge drop in standard reduction potential. The values
shown are approximate because the reduction potentials of the carriers vary with experimental conditions. The pathway shows the stoichiometry when
a pair of electrons is transferred from H 2 0 to NADPH. Abbreviations: Ph a, pheophytin a, electron acceptor of P680; PQ A , bound plastoquinone;

PQ b , mobile plastoquinone; A 0 , chlorophyll a, the primary electron acceptor of P700; A lf phyl loqu i none; F x , F B , and F A , iron-sulfur clusters; Fd,
ferredoxin; FNR, ferredoxin:NADP + reductase.

15.2 Bacterial Photosystems

457

The main difference between photosynthesis and respiratory electron transport is
the source of electrons and the terminal electron acceptors. In mitochondria, for exam-
ple, “high energy” electrons are supplied by reducing equivalents such as NADH (E of =
-0.32 V) and accepted by 0 2 ( E ° ' = +0.82 V) to produce water. In the coupled photo-
synthesis pathway the flow of electrons is reversed — water ( E°' = +0.82 V) is the
electron donor and NADP 0 (E°' = —0.32 V) is the electron acceptor. This “reversal” of
electron flow is thermodynamically unfavorable unless it is coupled to other reactions
with a larger Gibbs free energy change. Those other reactions are, of course, the excita-
tion of PSI and PSII by sunlight.

In order to extract electrons from water the cell needs to generate a powerful
oxidizing agent with a reduction potential greater than that of the H 2 0 1/2 0 2 +

2H© + 2e 0 reaction. This strong oxidizing agent is the P680 special pair after it has
given up an electron. The half reaction is P68O 0 + e 0 — » P680° {E°’ = +1.1 V). Note that
this standard reduction potential is higher than that of water so that electrons can flow
“down” from water to P68O 0 as shown in Figure 15.14. P68O 0 is the most powerful
oxidizing agent in biochemical reactions. It is much more potent than P87O 0 in purple
bacteria even though purple bacteria have a similar type II reaction center.

Similarly, P700* is a strong reducing agent with a lower reduction potential than
NADP 0 . In this case, the absorption of a photon of light by PSI creates an energetic
electron that can be passed “down” to NADP 0 to create reducing equivalents in the
form of NADPH. Thus, the “reversal” of electron flow in photosynthesis, compared to
respiratory electron transport, is achieved by the special light-absorbing properties of
chlorophyll molecules in the two photosystems.

E. Photosynthesis Takes Place Within Internal Membranes

All four of the photosynthesis complexes (PSI, PSII, cytochrome bfi and ATP synthase)
are embedded in membranes. Most cyanobacteria contain a complex internal network
of membranes where these complexes are concentrated (Figure 15.15). The internal
membranes are called thylakoid membranes. They form by invagination of the inner
plasma membrane creating structures that are similar to the mitochondrial cristae. As
the membrane folds inward it encloses a space called the lumen where protons accumu-
late during photosynthesis. The thylakoid lumen may remain connected to the periplas-
mic space or it may form an internal compartment if a membrane loop (or bubble)
pinches off from the plasma membrane.

Plasma Thylakoid

membrane membranes

Carboxysomes Peptidoglycan
layer

— 100 nm

▲ Figure 15.15

Internal structure of the cyanobacterium

Synechocystis PCC 6803. (Carboxysomes are
described in Section 15. 6A.)

BOX 15.2 OXYGEN “POLLUTION” OF EARTH’S ATMOSPHERE

Photosynthetic bacteria probably evolved three billion years
ago but the earliest fossil evidence of oxygen producing
cyanobacteria dates only from 2.1 billion years ago — claims
of much earlier fossils have recently been discredited. The ge-
ological record strongly indicates that bacteria began “pollut-
ing” the atmosphere with oxygen about 2.4-2. 7 billion years
ago. This likely corresponds to the evolution of the oxygen
evolving complex in PSII and it predates the earliest cyanobac-
teria fossils.

At that time, oxygen levels rose to about 25% of the pres-
ent level and they remained at that level for more than a billion
years except for a brief drop around 1.9 billion years ago. The
cause of this decline isn’t known. Primitive plants — probably
lichens and mosses — invaded land about 700 million years ago
and this led to a steep rise in oxygen levels that eventually
reached the present-day concentration of 21%.

Oxygen was highly toxic to most of the species that were
around 2 billion years ago but gradually new species arose
that could not only tolerate the “pollutant” but used it in res-
piratory electron transport.

▲ Oxygen levels in Earth’s atmosphere.

458 CHAPTER 15 Photosynthesis

▲ Chlamydomonas sp. Chlamydomonas
species are green algae that are closely re-
lated to plants. They contain a single large
chloroplast. “Chlamy” is a model organism
that is easily grown in the laboratory.

▲ Diatoms. About 30% of the oxygen in our
atmosphere comes from marine photosyn-
thetic organisms.

The internal membrane network presents a much greater surface area for mem-
brane proteins. As a result, cyanobacteria contain a much higher concentration of pho-
tosynthesis complexes compared to other species of photosynthetic bacteria. This
means that cyanobacteria are very efficient at capturing light energy and converting it to
chemical energy. This, in turn, has led to their evolutionary success and the formation
of an oxygen enriched atmosphere.

15.3 Plant Photosynthesis

Up to this point we have been describing bacterial photosynthesis but many eukaryotic
species are capable of photosynthesis. The photosynthesizing eukaryotes we are most
familiar with are flowering plants and other terrestrial species such as mosses and ferns.
In addition to these obvious examples, there are many simpler species such as algae and
diatoms.

In all photosynthesizing eukaryotes the light- gathering photosystems are localized
to a specific cellular organelle called the chloroplast. Thus, unlike bacterial metabolism,
photosynthesis and respiratory electron transport are not integrated since they take
place in different compartments (chloroplasts and mitochondria). Chloroplasts evolved
from a species of cyanobacteria that entered into a symbiotic relationship with a primi-
tive eukaryotic cell over 1 billion years ago. Modern chloroplasts still retain a reduced
form of the original bacterial genome. This DNA contains many of the genes for the
proteins of the photosystems and genes for some of the enzymes involved in C0 2 fixa-
tion. The transcription of these genes and the translation of their mRNAs resemble the
prokaryotic mechanisms described in Chapters 21 and 22. This prokaryotic flavor of
gene expression reflects the evolutionary origin of chloroplasts.

In the modern world, a large percentage (~70%) of total atmospheric oxygen is
produced by photosynthesis in land plants, especially in tropical rain forests. The re-
maining oxygen is produced by small marine organisms, mostly bacteria, diatoms, and
algae. Almost all of the food for animals comes directly or indirectly from plants and the
synthesis of these food molecules relies on the energy of sunlight.

A. Chloroplasts

The chloroplast is enclosed by a double membrane (Figure 15.16). As in mitochondria,
the outer membrane is exposed to the cytoplasm and the inner membrane forms highly
folded internal structures. During photosynthesis protons are translocated from the inte-
rior of the chloroplast, called the stroma, to the compartments between the membranes.

The interior membrane is called the thylakoid membrane. Recall that cyanobacteria
possess a similar thylakoid membrane (Figure 15.15). In the chloroplast this membrane
forms an extensive network of sheets within the organelle. As the chloroplast develops,
projections grow out from these sheets to form flattened disk-like structures. These disk-
like structures stack on top of one another like a pile of coins to form grana (singular,
granum). A typical chloroplast contains dozens of grana, or stacked disks of thylakoid
membranes. The grana in mature chloroplasts are connected to each other by thin sheets
of thylakoid membrane called stroma thylakoids. These stroma thylakoid membranes are
exposed to the stroma on both surfaces whereas grana thylakoid membranes within a
stack are in close contact with the membranes immediately above and below them.

The three-dimensional organization of the thylakoid membrane is shown in
Figure 15.17. Each disk in the stack is connected to the stroma thylakoids by short
bridges. The interior of each disk is called the lumen and it is the same compartment as
the region between the two membranes of the stroma thylakoid. All thylakoid mem-
branes are likely derived from the inner chloroplast membrane. This means that the
lumen is topologically equivalent to the space between the inner and outer membranes
of the chloroplast although in some cases the direct connection may be lost. The thy-
lakoid membranes contain PSI, PSII, cytochrome bf, and ATP synthase complexes as in
cyanobacteria. In mitochondria, protons accumulate in the compartment between the
inner and outer membranes (Section 14.3); similarly, in chloroplasts, protons are
translocated into the thylakoid lumen and the space between the two membranes of

15.3 Plant Photosynthesis 459

(a)

(b)

Intermembrane Outer

space membrane Inner

membrane

Stroma

Granum

Lumen
Stromal

Thylakoid Granal lamellae
membrane lamellae

▲ Figure 15.16

Structure of the chloroplast. (a) Illustration, (b) Electron micrograph: cross-section of a chloroplast from a spinach leaf. Shown are grana (G), the
thylakoid membrane (T), and the stroma (S).

the stroma thylakoids. It’s important to keep in mind that the chloroplast stroma is
equivalent to the cytoplasm in bacteria and the matrix in mitochondria.

B. Plant Photosystems

The photosynthesis complexes in eukaryotic chloroplasts evolved from the complexes
present in primitive cyanobacteria. Chloroplast PSI is structurally and functionally sim-
ilar to its bacterial ancestor — the only significant structural difference is that eukaryotic
PSI contains chlorophyll molecules instead of bacteriochlorophyll in the electron transfer
chain of the reaction center. The eukaryotic version oxidizes plastocyanin (or cytochrome c)
and reduces ferredoxin (or flavodoxin). Eukaryotic PSI associates with a light-
harvesting complex called LHCI that resembles the complex found in some bacteria.

Chloroplast PSII is also similar to the one in cyanobacteria. Plant chloroplasts
contain a light-harvesting complex called LHCII that associates with PSII in the
chloroplast membrane. LHCII is a large structure containing 140 chlorophylls and 40
carotenoids and it completely surrounds PSII. As a result, photon capture in plants is
more efficient than in bacteria. Cyanobacteria and chloroplasts contain similar cy-
tochrome bf complexes.

The ATP synthase in chloroplasts is related to the cyanobacterial ATP synthase, as
expected. The protein components differ from the mitochondrial version described in
the previous chapter. This is not surprising since the mitochondrial ATP synthase
evolved from the proteobacterial ancestor of bacteria and proteobacteria are distantly re-
lated to cyanobacteria. Species such as algae, diatoms, and plants that contain both mito-
chondria and chloroplasts have distinctive versions of ATP synthase in each organelle.

The chloroplast ATP synthase is a CFoFi ATPase where the cc C” stands for chloro-
plast. The overall molecular structure is very similar to that of mitochondria even
though the various subunits of the two enzymes are encoded by different genes. As in
mitochondria, the membrane component of the chloroplast ATP synthase consists of a
multimeric ring and a rod that projects into a hexameric head structure. The ring ro-
tates as protons move across the membrane and ATP is synthesized from ADP + Pj by a
binding change mechanism as described in Section 14.9. The “knob” projects into the
chloroplast stroma (Figure 15.18).

C. Organization of Chloroplast Photosystems

Figure 15.19 illustrates the locations of the membrane-spanning photosynthetic com-
ponents within the chloroplast thylakoid membrane. PSI is located predominantly in
the stroma thylakoid and is therefore exposed to the chloroplast stroma. PSII is located

▲ Figure 15.17

Organization of stacked disks in a granum and
their connection to the stroma thylakoids.

Adapted from Staehlin, L. A. (2003) Chloroplast
structure: from chlorophyll granules to supra-
molecular architecture of thylakoid membranes.
Photosyn thesis Research 76:185-196.

The locations of various photosynthetic
components in the stroma and grana
thylakoid membranes are shown in
Figure 15.19.

KEY CONCEPT

Photosynthetic bacteria and chloroplasts
make use of internal thylakoid membranes
to increase the number of photosystem
complexes.

460 CHAPTER 15 Photosynthesis

▲ Figure 15.18
Chloroplast ATP synthase.

predominantly in the grana thylakoid membrane, away from the stroma. The oxygen-
evolving complex is associated with PSII on the luminal side of the thylakoid mem-
brane. The cytochrome fr/ complex spans the thylakoid membrane and is found in both
the stroma and grana thylakoid membranes. ATP synthase is found exclusively in the
stroma thylakoids with the CVi component, the site of ATP synthesis, projecting into
the stroma.

The membranes of the top and bottom surfaces of each disk in a granum are in
contact with each other forming a double-membrane structure. This region is densely
packed with the light-absorbing PSII complexes and their associated LHCII complexes.
Light passes through the plasma membrane of the plant cell, through the cytoplasm,
and across the outer membrane of the chloroplast. When light reaches the grana, the
photons are efficiently absorbed by the pigment molecules in the membrane.

Excited electrons are transferred within PSII to PQ forming PQH 2 . The protons
for this reaction are taken up from the stroma. The PSII reaction center is replenished
with electrons from the oxidation of water taking place in the lumen. PQH 2 diffuses
within the membrane to the cytochrome fr/ complex where it is oxidized to PQ. The
protons released in the Q cycle enter the lumen. Electrons are passed to plastocyanin
that diffuses freely in the lumen to reach PSI. PSI absorbs light leading to the transfer of
electrons from reduced plastocyanin to ferredoxin. Ferredoxin is formed in the stroma.
It can participate in the reduction of NADP® to NADPH in the stroma or serve as an
electron donor to cytochrome bf complexes in the stroma thylakoid membrane (cyclic
electron transport, Section 15.2B).

Note that PSII is not directly exposed to the stroma but is exposed to the thylakoid
lumen. The lumen is topologically equivalent to the outside of the bacterial membrane
as shown in Figure 15.11. PSI projects into the stroma compartment since it produces
ferredoxin that accumulates within chloroplasts. The stroma is topologically equivalent
to the bacterial cytoplasm (inside the cell). The distribution of cytochrome fr/ com-
plexes is explained by the fact they can receive electrons from both PSII and PSI. Super-
complexes of PSII and cytochrome bf in the grana participate in linear electron transfer
from water to plastocyanin. In the stroma thylakoids there are complexes of PSI, cy-
tochrome bf and ferredoximquinone oxidoreductase (FQR) that are involved in cyclic
electron flow.

The proton gradient is used to generate ATP. As protons are translocated from the
lumen compartment to the stroma, ATP is synthesized from ADP and Pj in the stroma.
Both ATP and NADPH accumulate in the stroma where they can be used in biosynthesis
reactions. In plants, but not other photosynthetic species, a high percentage of ATP and
NADPH molecules are used in the fixation of C0 2 and the synthesis of carbohydrates.

Figure 15.19 ►

Distribution of membrane-spanning photosyn-
thetic components between stroma and granal
thylakoids. PSI is found predominantly in
stroma thylakoids. PSII is found predomi-
nantly in grana thylakoids. The cytochrome
bf complex in found in both stroma and
grana thylakoid membranes. ATP synthase is
localized exclusively to stroma thylakoids.

Photosystem II
Photosystem I
ATP synthase
Cyt bf complex

i j§

Grana

thylakoid

STROMA

Stroma

thylakoid

15.4 Fixation of CO 2 : The Calvin Cycle 461

BOX 15.3 BACTERIORHODOPSIN

Bacteriorhodopsin is a membrane protein found in a few
specialized species of archaebacteria such as Halobacterium
salinarium. The protein has seven membrane-spanning a
helices that form a channel in the membrane. (See ribbon
structure below.) A single retinal molecule is covalently
bound to a lysine side chain in the middle of the channel.
The normal configuration of the retinal is all- trans but when
it absorbs a photon of light it converts to the 13-ds configu-
ration. (See structure below.) The light-induced change in
configuration is coupled to deprotonation and reprotonation
of the retinol molecule.

When light is absorbed, the shift in configuration to
13-ds retinal releases a proton that then passes up the channel
to be released on the outside of the membrane. This proton is
replaced by a proton that is taken up from the cytosol and the
retinol configuration shifts back to the all - trans form. For
every photon of light that is absorbed by bacteriorhodopsin a
single proton is translocated across the membrane.

Bacteriorhodopsin creates a light- induced proton gradi-
ent and this proton gradient drives ATP synthesis by ATP
synthase.

INSIDE

a Bacteriorhodopsin.

OUTSIDE

- Membrane

a Two configurations of retinal-lysine in bacteriorhodopsin. (a) AW- trans
retinal, (b) 13-ds retinal. The configuration shifts from the all -trans
form to the 13-dsform when a photon of light is absorbed.

The coupling of bacteriorhodopsin and ATP synthase
can be directly demonstrated by artificially synthesizing lipid
vesicles containing both complexes. In the orientation shown
below, the vesicles will synthesize ATP from ADP + Pj when
they are illuminated. This experiment, first carried out by
Efraim Racker and his colleagues in 1974, was one of the first
confirmations of the chemiosmotic theory (Section 14.3).

Bacteriorhodopsin

a Bacteriorhodopsin creates a proton gradient that drives ATP synthesis.

Artificial lipid vesicles containing bacteriorhodopsin and ATP syn-
thase were created with the orientation shown. When these vesicles
were illuminated, bacteriorhodopsin pumped protons into the vesicle
and the resulting proton gradient activated ATP synthase.

15.4 Fixation of CO 2 : The Calvin Cycle

In photosynthetic species there is a special pathway for the reductive conversion of at-
mospheric C0 2 to carbohydrates. The reactions are powered by the ATP and NADPH
formed during the light reactions of photosynthesis. The fixation of C0 2 and the syn-
thesis of carbohydrates occurs in the cytoplasm of bacteria and in the chloroplast
stroma. This biosynthesis pathway is a cycle of enzyme -catalyzed reactions with three
major stages: (1) the carboxylation of a five-carbon sugar molecule, (2) the reductive
synthesis of carbohydrate for use in other pathways, and (3) the regeneration of the
molecule that accepts C0 2 . This pathway of carbon assimilation has several names, such

462 CHAPTER 15 Photosynthesis

▲ Melvin Calvin (191 1-1997). Calvin won the
Nobel Prize in Chemistry in 1961 for his
work on carbon dioxide assimilation in plants.

Ibl.gov/Science-Articles/Research-Review/Magazine/

1997/storyl2.html]

KEY CONCEPT

The Calvin cycle utilizes the products of
photosynthesis, ATP and NADPH, to fix
C0 2 into carbohydrates.

as the reductive pentose phosphate cycle , the C 3 pathway (the first intermediate is a three-
carbon molecule), and the Calvin cycle. (Workers in Melvin Calvins laboratory discov-
ered the carbon-fixing pathway using 14 C0 2 tracer experiments in algae.) We refer to
the pathway as the Calvin cycle.

The fixation of C0 2 and the synthesis of carbohydrates are often described as “pho-
tosynthesis.” In this textbook we refer to photosynthesis and the Calvin cycle as two
separate pathways.

A. The Calvin Cycle

The Calvin cycle is outlined in Figure 15.20. The first stage is the carboxylation of ribu-
lose 1,5-frzsphosphate, a reaction catalyzed by the enzyme ribulose 1,5-frzsphosphate
carboxylase-oxygenase, better known as Rubisco. The second stage is a reduction stage
where 3-phosphoglycerate is converted to glyceraldehyde 3 -phosphate. Most of the
glyceraldehyde 3-phosphate is converted to ribulose 1,5-frisphosphate in the third (re-
generation) stage. Some of the glyceraldehyde 3 -phosphate produced in the Calvin cycle
is used in carbohydrate synthesis pathways. Glyceraldehye 3-phosphate is the main
product of the Calvin cycle.

Figure 15.21 on page 464 shows all reactions of the Calvin cycle. The pathway be-
gins with steps for assimilating three molecules of carbon dioxide because the smallest
carbon intermediate in the Calvin cycle is a C 3 molecule. Therefore, three C0 2 mole-
cules must be fixed before one C 3 unit (glyceraldehyde 3 -phosphate) can be removed
from the cycle without diminishing the metabolic pools.

B. Rubisco: Ribulose 1 ,5-6/sphosphate Carboxylase-oxygenase

Rubisco (ribulose l,5-Z?isphosphate carboxylase-oxygenase) is the key enzyme of the
Calvin cycle. It catalyzes the fixation of atmospheric C0 2 into carbon compounds.
This reaction involves the carboxylation of the five-carbon sugar, ribulose 1,5-
frisphosphate, by C0 2 . This leads to the eventual release of two three-carbon molecules
of 3-phosphoglycerate. The reaction mechanism of Rubisco is shown in Figure 15.22.

Rubisco makes up about 50% of the soluble protein in plant leaves, making it one of
the most abundant enzymes on Earth. Interestingly, its status as an abundant enzyme is
due partly to the fact that it is not very efficient — the low turnover number of ~3 s -1
means that large amounts of the enzyme are required to support C0 2 fixation!

The Rubisco of plants, algae, and cyanobacteria is composed of eight large (L) sub-
units and eight small (S) subunits (Figure 15.23). There are eight active sites located in
the eight large subunits. Four additional small subunits are located at each end of the
core formed by the large subunits. The Rubisco molecules in other photosynthetic bac-
teria have only the large subunits containing the active sites. For example, in the purple
bacterium Rho do spirillum rubrum , Rubisco consists of a simple dimer of large subunits.

Figure 15.20 ►

Summary of the Calvin cycle. The cycle has
three stages: carboxylation of ribulose 1,5-
b/'sphosphate, reduction of 3-phosphoglycerate
to glyceraldehyde 3-phosphate, and regener-
ation of ribulose 1,5-b/sphosphate.

Ribulose C0 2

r 1,5-b/sphosphate

Carboxylation

^ , , , . f Regeneration

Carbohydrates^/ ^

Glyceraldehyde Reduction

3-Phosphoglycerate
-ATP

3-phosphate

ADP

1,3-£/sphosphoglycerate

Pi i NADPH

NADP® + H®

15.4 Fixation of CO 2 : The Calvin Cycle 463

ch 2 opo 3 ®

1 r\

c =0

r 1

H — C— OH

H — C— OH
CH 2 OP0 3 ®
Ribulose

1,5-b/sphosphate

j©

Enolization

j®

ch 2 opo 3 ©

c— O 0
II

C— OH

H — C— OH
CH,OPO,®

CO,

ch 2 opo 3 ©

C — O 0

C ^ H
H — C — OH

CH-.OPO

2,3-Enediolate

intermediate

Carboxylation

-l-UO

ch 2 opo 3

HO— c — H

' P

Protonation

coa

3-Phosphoglycerate

j®

ch 2 opo 3 ®

HO— C

©\ 0

coo u

Carbanion

ch 2 opo 3 ©

CH 2 0P0 3 ®

Cleavage

1 (P)

HO — C— COO 0

1 q

HO— C — COO u u

1 /

x —

2 H®

HO — C -rO-p H
1 ^

b 0

(°2
-u —

H — C— OH

coo°

ch 2 opo 3 ©

— c

:— oh

Gem diol

2-Carboxy-3-ketoarabinitol

ch 2 opo 3 ®

3-Phosphoglycerate

intermediate

1,5-b/sphosphate

▲ Figure 15.22

Mechanism of Rubisco-catalyzed carboxylation of ribulose 1 ,5-Z;/sphosphate to form two molecules of 3-phosphoglycerate. A proton is abstracted from C-3
of ribulose 1,5-b/sphosphate to create a 2,3-enediolate intermediate. The nucleophilic enediolate attacks C0 2 , producing 2-carboxy-3-ketoarabinitol
1,5-b/sphosphate, which is hydrated to an unstable gem diol intermediate. The C-2-C-3 bond of the intermediate is immediately cleaved, generating
a carbanion and one molecule of 3-phosphoglycerate. Stereospecific protonation of the carbanion yields a second molecule of 3-phosphoglycerate.
This step completes the carbon fixation stage of the RPP cycle — two molecules of 3-phosphoglycerate are formed from C0 2 and the five-carbon sugar
ribulose 1,5-b/sphosphate.

The purple bacterium version of Rubisco has a much lower affinity for C0 2 than
the more complex multisubunit enzymes in other species but it catalyzes the same reac-
tion. In a spectacular demonstration of this functional similarity, tobacco plants were
genetically engineered by replacing the normal plant gene with the one from the purple
bacterium Rho do spirillum rubrum. The modified plants contained only the dimeric
bacterial form of the enzyme but they grew normally and reproduced as long as they
were kept in an atmosphere of high C0 2 concentration.

(b)

◄ Figure 15.23

The quaternary structure (L 8 S 8 ) of ribulose
1 ,5-b/sphosphate carboxylase-oxygenase
(Rubisco). (a) Top and (b) side views of the
enzyme from spinach ( Spinacia oleracea).
Large subunits are shown alternately yellow
and blue; small subunits are purple.

[PDB 1RCX].

+->

£ o

(D
+->
<D 03
to (—

O Q.

_L_

C£ Q.

▲ Figure 15.21

Calvin cycle. The concentrations of Calvin cycle intermediates are maintained when one molecule of glyceraldehyde 3-phosphate (G3P) exits the
cycle after three molecules of C0 2 are fixed.

464

15.4 Fixation of CO2: The Calvin Cycle 465

Rubisco cycles between an active form (in the light) and an inactive form (in the
dark). It must be activated to catalyze the fixation of C0 2 . In the light, Rubisco activity
increases in response to the higher, more basic pH that develops in the stroma (or bac-
terial cytoplasm) during proton translocation. Under alkaline conditions an activating
molecule of C0 2 , which is not the substrate C0 2 molecule, reacts reversibly with the
side chain of a lysine residue of Rubisco to form a carbamate adduct. Mg© binds to and
stabilizes this C0 2 -lysine adduct. The enzyme must be carbamylated in order to carry
out C0 2 fixation; however, the carbamate adduct readily dissociates, making the enzyme
inactive. Carbamylation is normally inhibited because Rubico is usually in an inactive
conformation. During the day, a light- activated ATP-dependent enzyme called Rubisco
activase binds to Rubisco and facilitates carbamylation by inducing a conformational
change. Under these conditions Rubisco is active.

When the sun goes down Rubisco activase is no longer effective in activating Ru-
bisco and C0 2 fixation stops. This regulation makes sense since photosynthesis is not
active at night and ATP + NADPH are not produced in chloroplasts during the night.
These cofactors are required for the Calvin cycle so the Calvin cycle is not active at night
as a result of the regulation of Rubisco activity. Inhibition of Rubisco in the dark pre-
vents the inefficient accumulation of 3-phosphoglycerate and the wasteful oxygenation
reaction described in the next section.

In plants, an additional level of inhibition is mediated by 2-carboxyarabinitol 1-
phosphate (Figure 15.24). This compound is an analog of the unstable gem diol inter-
mediate of the carboxylation reaction. It is synthesized only at night and it binds to, and
inhibits, any residual carbamylated Rubisco, thus ensuring that the Calvin cycle is shut
down. Some plants synthesize sufficient amounts of the inhibitor to keep Rubisco com-
pletely inactive in the dark.

ch 2 opo 3 ®

HO — c— COO 0

H — C— OH

ch 2 oh

▲ Figure 15.24

2-Carboxyarabinitol 1 -phosphate.

C. Oxygenation of Ribulose 1 ,5-6/sphosphate

As its complete name indicates, ribulose 1,5-Hsphosphate carboxylase-oxygenase cat-
alyzes not only carboxylation but also the oxygenation of ribulose 1,5-frzsphosphate.
The two reactions are competitive since C0 2 and 0 2 compete for the same active sites
on Rubisco. The oxygenation reaction produces one molecule of 3-phosphoglycerate
and one molecule of 2-phosphoglycolate (Figure 15.25). Oxygenation consumes signif-
icant amounts of ribulose 1,5-frzsphosphate in vivo. Under normal growth conditions,
the rate of carboxylation is only about three to four times the rate of oxygenation.

The 3-phosphoglycerate formed from the oxygenation of ribulose 1,5-
frisphosphate enters the Calvin cycle. The other product of the oxygenation reaction fol-
lows a different pathway. Two molecules of 2-phosphoglycolate (C 2 ) are metabolized in
peroxisomes and the mitochondria by an oxidative pathway (via glyoxylate and the
amino acids glycine and serine) to one molecule of C0 2 and one molecule of 3-
phosphoglycerate (C 3 ), which also enters the Calvin cycle. This oxidative pathway con-
sumes NADH and ATR The light-dependent uptake of 0 2 catalyzed by Rubisco and fol-
lowed by the release of C0 2 during the metabolism of 2-phosphoglycolate is called
photorespiration. Like carboxylation, photorespiration is normally inhibited in darkness
when Rubisco is inactive. The appreciable release of fixed C0 2 and the consumption of

KEY CONCEPT

Some enzymes cannot distinguish
between very similar substrates.

ch 2 opo 3 ®

C =0

H — C— OH

CH 2 OP0 3 ©

Ribulose

1,5-b/sphosphate

ch 2 opo 3 ®

coo 0

2-Phosphoglycolate

coo 0

H — C — OH

ch 2 opo 3 ©

3-Phosphoglycerate

◄ Figure 15.25

Oxygenation of ribulose 1 ,5-ib/sphosphate
catalyzed by Rubisco.

466 CHAPTER 15 Photosynthesis

BOX 15.4 BUILDING A BETTER RUBISCO

Many labs are attempting to genetically modify plants in
order to enhance the carboxylation reaction and suppress the
oxygenation reaction. If successful, these attempts to make a
better Rubisco could greatly increase food production.

The “perfect” enzyme would have very low oxygenase ac-
tivity and very efficient carboxylase activity. The kinetic param-
eters of the oxygenase activity of Rubisco enzymes from several
species are listed in the accompanying table. The low catalytic
efficiency of the enzyme is indicated by the k Cdit /K m values.

Kinetic parameters of Rubisco carboxylase activity in various species

Species

*cat (S” 1 )

0*M)

(M-V 1 )

Tobacco

3.4

10.7

3.2 X 10 5

Red algae

2.6

9.3

2.8 X 10 5

Purple bacteria

7.3

8.2 X 1 0 4

"Perfect" enzyme

1070

10.7

10 8

Data from Andrews, J. T., and Whitney, S. M. (2003). Manipulating ribulose
b/sphosphate carboxylase/oxygenase in the chloroplasts of higher plants. Arch.
Biochem. Biophys. 414: 159-169.

These values should be compared to those in Table 5.2. It
seems likely that the carboxylase efficiency can be improved
1000-fold by modifying the amino acid side chains in the
active site.

The difficult part of the genetic modification is choosing
the appropriate amino acid changes. The choice is informed
by a detailed knowledge of the structures of several Rubisco
enzymes from different species and by examination of the
contacts between amino acid side chains and substrate mole-
cules. Models of the presumed transition states are also
important. Additional key residues can be identified by com-
paring the conservation of amino acid sequences in enzymes
from a wide variety of species

The underlying strategy assumes that evolution has not
yet selected for the most well-designed enzyme. This as-
sumption seems reasonable since there are many examples of
ongoing evolution in biochemistry. However, several billion
years of evolution have not resulted in a better Rubisco and
neither have several decades of human effort. It may not be
possible to build a better Rubisco.

3C 3C 3C 3C 3C

▲ Figure 15.26

Outline of the regeneration stage of the Calvin
cycle.

energy as a result of oxygenation — with no apparent benefit to the organism — arise
from the lack of absolute substrate specificity of Rubisco. This is a serious problem in
agriculture because photorespiration limits crop yields.

D. Calvin Cycle: Reduction and Regeneration Stages

The reduction stage of the Calvin cycle begins with the ATP-dependent conversion of
3-phosphoglycerate to 1,3-frisphosphoglycerate in a reaction catalyzed by phosphoglycer-
ate kinase. Next, 1,3-frisphosphoglycerate is reduced by NADPH (not NADH, as in glu-
coneogenesis, Section 11.2#6) in a reaction catalyzed by a glyceraldehyde 3-phosphate
dehydrogenase isozyme. As in gluconeogenesis, some of the glyceraldehyde 3 -phosphate
is rearranged to its isomer, dihydroxyacetone phosphate, by triose phosphate isomerase.
For every six glyceraldehyde 3 -phosphate molecules produced by this pathway, one is
removed from the cycle to be used in carbohydrate synthesis and the five others are used
in the regeneration stage.

In the regeneration stage, glyceraldehyde 3 -phosphate is diverted into three different
branches of the pathway and is interconverted between three-carbon (3C), four-carbon
(4C), five-carbon (5C), six-carbon (6C), and seven-carbon (7C) phosphorylated sugars
(Figure 15.21). The pathway is schematically outlined in Figure 15.26. Two of the reactions,
those catalyzed by aldolase and fructose 1.6-Hsphosphatase, are familiar because they
are part of the gluconeogenesis pathway (Section 12.1). Many of the other reactions are
part of the normal pentose phosphate pathway (Section 12.4) including two tranketolase
reactions. The net result of the Calvin cycle reactions is

3 C0 2 + 9 ATP + 6 NADPH + 5 H 2 0 >

glyceraldehyde 3-phosphate + 9 ADP + 8 Pj + 6 NADP© + 2H + (15.2)

Both ATP and NADPH are required for C0 2 fixation by the Calvin cycle. These are the
major products of the light reactions of photosynthesis. The fact that the requirement
for ATP exceeds that of NADPH is one reason why cyclic electron flow from PSI to cy-
tochrome bf is important in photosynthesis. Cyclic electron flow results in increased
production of ATP relative to NADPH.

15.5 Sucrose and Starch Metabolism in Plants 467

It’s interesting to compare the cost of synthesizing carbohydrates from C0 2 and the
energy yield from degrading it via glycolysis and the citric acid cycle. We can use Reaction
15.2 to estimate the cost of synthesizing acetyl CoA — the substrate for the citric acid
cycle. Recall that the pathway from glyceraldehyde 3 -phosphate to acetyl CoA is coupled
to the synthesis of two molecules of NADH and two molecules of ATP (Section 1 1.2). If
we subtract these from the cost of making glyceraldehyde 3 -phosphate then the total
cost of synthesizing acetyl CoA from C0 2 is 7 ATP + 4 NAD(P)H. This can be expressed
as 17 ATP equivalents since each NADH is equivalent to 2.5 ATP (Section 14.11). The
net gain from complete oxidation of acetyl CoA by the citric acid cycle is 10 ATP
equivalents (Section 13.4). The biosynthesis pathway is more expensive than the energy
gained from catabolism. In this case, the “efficiency” of acetyl CoA oxidation is only
about 60% (10/17 = 59%) but this value is misleading since it’s actually the biosynthesis
pathway (costing 17 ATP equivalents) that is complex and inefficient.

We can estimate the cost of synthesizing glucose because it is simply the cost of
making two molecules of glyceraldehyde 3 -phosphate. It’s equivalent to 18 molecules of
ATP and 12 molecules of NADPH or 48 ATP equivalents. Recall that the net gain of en-
ergy from the complete oxidation of glucose via glycolysis and the citric acid cycle is 32
ATP equivalents (Section 13.4). In this case, catabolism recovers two-thirds of amount
of the ATP equivalents used in the biosynthesis pathway.

▲ Glyceraldehyde 3-phosphate dehydrogenase.

This NADPH-dependent enzyme from
spinach ( Spinacia oleracea) crystallizes as a
tetramer. Only a single subunit is shown
here. NADPH is bound in the active site of
the enzyme. [PDB 2PKQ]

15.5 Sucrose and Starch Metabolism in Plants

Glyceraldehyde 3 -phosphate (G3P) is the main product of carbon fixation in most pho-
tosynthetic species. G3P is subsequently converted to glucose by the gluconeogenesis
pathway. Newly synthesized hexoses can be used immediately as substrates in a number
of biosynthesis pathways or they can be stored as polysaccharides for use later on. In
bacteria, most algae, and some plants, the storage polysaccharide is glycogen, just as in
animals. The storage polysaccharide in vascular plants is usually starch.

Starch is synthesized in chloroplasts from glucose 6-phosphate, the primary product
of gluconeogenesis (Section 12. ID). In the first step, glucose 6-phosphate is converted
to glucose 1 -phosphate in a reaction catalyzed by phosphoglucomutase (Figure 15.27).
This is the same enzyme we encountered in the glycogen synthesis pathway (Section
12.5A). The second step is the activation of glucose by synthesis of ADP-glucose. This
reaction is catalyzed by ADP-glucose pyrophosphorylase. The metabolic strategy is
similar to that of glycogen biosynthesis except that the key intermediate in glycogen

KEY CONCEPT

The energy recovered in catabolic
pathways is usually about two-thirds of
the energy used in biosynthesis.

The structures of starch and glycogen
are described in Section 8.6A.

The nucleotide sugar ADP-glucose is
also required for synthesis of glycogen
by some bacteria (Section 12.5A).

468 CHAPTER 15 Photosynthesis

▲ Maple syrup. The sucrose-rich sap of
maple trees is collected and concentrated to
produce maple syrup.

synthesis is UDP-glucose. The polymerization reaction in starch biosynthesis is carried
out by starch synthase. This pathway consumes one molecule of ATP and releases one
molecule of pyrophosphate for each residue that is added to the growing polysaccharide
chain. ATP is supplied by the reactions of photosynthesis.

Starch is synthesized in daylight when photosynthesis is active and ATP molecules
accumulate within the chloroplast. During the night starch becomes a source of carbon
and energy for the plant. The starch molecule is cleaved by the action of starch phos-
phorylase to generate glucose 1 -phosphate that is converted to triose phosphates by
glycolysis. The triose phosphates are exported from the chloroplast to the cytoplasm.
Alternatively, starch can be hydrolyzed by the action of amylases to dextrins and eventu-
ally to maltose and then glucose. Glucose formed via this route is phosphorylated by the
action of hexokinase and enters the glycolytic pathway.

Sucrose is a mobile form of carbohydrate in plants. It is synthesized in the cyto-
plasm of cells that contain chloroplasts (e.g., leaf cells) and exported to the plant vascular
system where it is taken up by non- photosynthetic cells (e.g., root cells). Thus, sucrose is
functionally equivalent to glucose, the mobile form of carbohydrate in those animals
that possess a circulatory system (Section 12.5).

The pathway for sucrose synthesis is shown in Figure 15.28. Four molecules of
triose phosphate produce one molecule of sucrose. The triose phosphates follow the
gluconeogenesis pathway, condensing to form fructose 1,6-frisphosphate that is hydrolyzed
to yield fructose 6-phosphate. Fructose 6-phosphate isomerizes to glucose 6-phosphate
that is diverted from the gluconeogenesis pathway and converted to a-D-glucose
1 -phosphate. Glucose 1 -phosphate reacts with UTP to form UDP-glucose and this activated
glucose molecule donates its glucosyl group to a molecule of fructose 6-phosphate,

(2) Glyceraldehyde
3-phosphate

Aldolase

(2) Dihydroxyacetone
phosphate <-

(2) Fructose
1,6-b/sphosphate

Sucrose

phosphate

phosphatase

Glucose

6-phosphate

isomerase

Fructose (2) Fructose 6-phosphate

Glucose 6-phosphate

Phospho-

glucomutase

Sucrose Sucrose 6-phosphate

(u-D-Glucopyranosyl-
/3-D-fructofuranoside)

▲ Figure 15.28

Biosynthesis of sucrose from glyceraldehyde 3-phosphate and dihydroxyacetone phosphate in the cytosol. Four molecules of triose phosphate (4 C 3 ) are

converted to one molecule of sucrose (C 12 ).

15.6 Additional Carbon Fixation Pathways 469

BOX 15.5 GREGOR MENDEL’S WRINKLED PEAS

One of the genetic traits that Gregor Mendel studied was round (R) vs. wrinkled (r)
peas. The wrinkled pea phenotype is caused by a defect in the gene for starch
branching enzyme. Starch synthesis is partially blocked in the absence of this en-
zyme and the developing peas have a higher concentration of sucrose. This causes
them to absorb more water than the normal peas and they swell to a larger size.
When the seeds begin to dry out the mutant peas lose more water and their outer
surface takes on a wrinkled appearance.

The mutation is caused by insertion of a transposon into the gene. It is a
recessive loss-of-function mutation because a single copy of the normal wild-type

Til T7^r

allele in heterozygotes can produce enough starch branching enzyme to produce
starch granules.

▲ Round and wrinkled peas in a pod.

to form sucrose 6-phosphate. The final step is the hydrolysis of sucrose 6-phosphate to
form sucrose.

Inorganic phosphate (Pj) is produced in the sucrose synthesis pathway by the reac-
tions catalyzed by fructose 1,6-frzsphosphatase and sucrose phosphate phosphatase.
Pyrophosphate (PPj) is produced in the reaction catalyzed by UDP-glucose pyrophos-
phorylase. The pathway consumes one ATP equivalent (as UTP). Sucrose synthesis and
glycogen synthesis require an activated glucose molecule in the form of UDP-glucose
whereas starch biosynthesis uses ADP-glucose.

The first metabolically irreversible step in the sucrose biosynthesis pathway is the
hydrolysis of fructose 1,6-frzsphosphate to yield fructose 6-phosphate and P^. The activ-
ity of fructose 1,6-frzsphosphatase is inhibited by the allosteric modulator fructose
2,6-frzsphosphate (Figure 12.9) — a molecule we encountered in our examinations of
glycolysis and gluconeogenesis. In plants, the level of fructose 2,6-Hsphosphate is regu-
lated by several metabolites that reflect the suitability of conditions for sucrose synthesis.

Sucrose is taken up by non-photosynthetic cells where it is degraded by sucrase (in-
vertase) to glucose and fructose that supply energy via glycolysis and the citric acid cycle
(Section 1 1.6A). These hexoses can also be converted to starch in those tissues that store
carbohydrate for future use. In root cells, for example, sucrose is converted to hexose
monomers and these sugars are taken up by specialized organelles called amyloplasts.
Amyloplasts are modified chloroplasts that lack the photosynthesis complexes but
retain the enzymes for starch synthesis. In some plants, such as potatos, turnips, and
carrots, the root cells can store huge reservoirs of starch.

15.6 Additional Carbon Fixation Pathways

As mentioned earlier, one of the most important problems with carbon fixation is the
inefficiency of Rubisco, especially the oxygenation reaction that greatly limits crop
yields (Section 15.4C) . Different species have evolved a variety of ways of overcoming
this problem.

A. Compartmentalization in Bacteria

Bacteria avoid the problems of photorespiration by confining Rubisco to specialized
compartments called carboxysomes. Carboxysomes are surrounded by a protein coat
that is impermeable to oxygen. Rubisco is localized to carboxysomes and so is the en-
zyme carbonic anhydrase that converts bicarbonate (HC0 3 ^) to C0 2 (see Section 2.10
and Figure 7.1). The advantage of compartmentalization is that Rubisco is supplied
with an abundant source of C0 2 while protecting it against 0 2 , thus avoiding the ineffi-
ciencies of photorespiration.

B. The C 4 Pathway

Several plant species avoid wasteful photorespiration by means of secondary pathways
for carbon fixation. The net effect of these secondary pathways is to increase the local

▲ Amyloplasts in potato cells.

▲ Potatoes are an excellent source of starch.

French fries are served in Quebec with gravy
and cheese curds. The dish is called poutine.

470 CHAPTER 15 Photosynthesis

▲ Carboxysomes.

Cyanobacteria ( Synechococcus elongatus )
cells are stained with a fluorescent dye
showing thylakoid membranes (red) and
carboxysomes (green).

Figure 15.29 ►

C 4 pathway. C0 2 is hydrated to bicarbonate
(HC0 3 “) in the mesophyll cytosol. Bicarbon-
ate reacts with phosphoenolpyruvate in a
carboxylation reaction catalyzed by phospho-
enolpyruvate (PEP) carboxylase, a cytosolic
enzyme that has no oxygenase activity.
Depending on the species, the oxaloacetate
produced is either reduced or transaminated
to form a four-carbon carboxylic acid or amino
acid, which is transported to an adjacent
bundle sheath cell and decarboxylated. The
released C0 2 is fixed by the Rubisco reaction
and enters the RPP cycle. The remaining
three-carbon compound is converted back
to the C0 2 acceptor, phosphoenolpyruvate.

concentration of C0 2 relative to 0 2 in those cells where Rubisco is active. One of these
pathways is called the C 4 pathway because it involves four-carbon intermediates. C 4 plants
tend to grow at high temperatures and high light intensities. They include such economi-
cally important species as maize (corn), sorghum, and sugarcane, and many of the most
troublesome weeds. The avoidance of photorespiration by tropical plants is essential
because the ratio of oxygenation to carboxylation by Rubisco increases with temperature.

The C 4 pathway concentrates C0 2 and delivers it to cells in the interior of the leaf
where the Calvin cycle is active. The initial product of carbon fixation is a four-carbon
acid (C 4 ) rather than a three-carbon acid as in the Calvin cycle. The C 4 pathway occurs
in two different cell types within the leaf. First, C0 2 is hydrated to bicarbonate that re-
acts with the C 3 compound phosphoenolpyruvate to form a C 4 acid in mesophyll cells
(near the leaf exterior). This reaction is catalyzed by an isozyme of phosphoenolpyru-
vate (PEP) carboxylase (Section 13.6). Next, the C 4 acid is transported to bundle sheath
cells in the interior of the leaf where it is decarboxylated. Because they are not directly
exposed to the atmosphere, the bundle sheath cells have a much lower 0 2 concentration
than mesophyll cells. The released C0 2 is fixed by the action of Rubisco and incorpo-
rated into the Calvin cycle. Phosphoenolpyruvate is regenerated from the remaining C 3
product. Figure 15.29 outlines the sequence of C 4 pathway reactions.

Atmospheric C0 2

15.6 Additional Carbon Fixation Pathways 471

The cell walls of internal bundle sheath cells are impermeable to gases. The decar-
boxylation of C 4 acids in these cells greatly increases the C0 2 concentration and creates
a high ratio of C0 2 to 0 2 . The oxygenase activity of Rubisco is minimized because there
is an insignificant amount of Rubisco in mesophyll cells and the ratio of C0 2 to 0 2 is
extremely high in bundle sheath cells. As a result, C 4 plants have essentially no pho-
torespiration activity. Although there is an extra energy cost to form phosphoenolpyru-
vate for C 4 carbon assimilation, the absence of photorespiration gives C 4 plants a signif-
icant advantage over C 3 plants.

C. Crassulacean Acid Metabolism (CAM)

Succulent plants, such as many species of cactus, grow primarily in arid environ-
ments where water loss can be a serious problem. A large amount of water can be
lost from the leaf tissues during carbon fixation since the cells must be exposed to
atmospheric C0 2 and water can evaporate from the surface. These plants minimize
water loss during photosynthesis by assimilating carbon at night. The pathway is
called Crassulacean acid metabolism because it was first discovered in the family
Crassulaceae.

The surface of the leaf in terrestrial vascular plants is often covered with an imper-
meable waxy coating and C0 2 passes through structures called stomata to reach pho-
tosynthetic cells. Stomata are formed by two adjacent cells on the surface of the leaf.
These guard cells define the entrance to a cavity lined with cells containing chloro-
plasts. The aperture between the guard cells changes in response to ion fluxes and the
resulting osmotic uptake of water. The flux of ions across the guard cells is regulated by
conditions that affect C0 2 fixation such as temperature and the availability of water. In
the heat of the day, CAM plants keep their stomata closed to minimize water loss. At
night, mesophyll cells take up C0 2 through open stomata. Water loss through the
stomata is much lower at cooler nighttime temperatures than during the day. C0 2 is
fixed by the PEP carboxylase reaction, and the oxaloacetate formed is reduced to
malate (Figure 15.30).

Malate is stored in a large central vacuole in order to maintain a nearly neutral pH
in the cytosol since the cellular concentration of this acid can reach 0.2 M by the end of
the night. The vacuoles of CAM plants generally occupy more than 90% of the total vol-
ume of the cell. Malate is released from the vacuole and decarboxylated during the day
when ATP and NADPH are formed by photosynthesis. Thus, the large pool of malate
accumulated at night supplies C0 2 for carbon assimilation during the day. Leaf stomata
are tightly closed when malate is decarboxylated so that neither water nor C0 2 can es-
cape from the leaf and the level of cellular C0 2 can be much higher than the level of at-
mospheric C0 2 . As in C 4 plants, the higher internal C0 2 concentration greatly reduces
photorespiration.

In CAM plants the phosphoenolpyruvate required for malate formation is derived
from starch via glycolysis. The phosphoenolpyruvate formed by malate decarboxyla-
tion (either directly by PEP carboxykinase or via malic enzyme and pyruvate
phosphate dikinase) is converted to starch via gluconeogenesis and stored in the
chloroplast.

CAM is analogous to C 4 metabolism in that the C 4 acid formed by the action of
PEP carboxylase is subsequently decarboxylated to supply C0 2 to the Calvin cycle. In
the C 4 pathway the carboxylation and decarboxylation phases of the cycle are spatially
separated in distinct cell types whereas in CAM they are temporally separated in day
and night cycles.

An important regulatory feature of the CAM pathway is the inhibition of PEP car-
boxylase by malate and low pH. PEP carboxylase is effectively inhibited during the day
when the cytosolic concentration of malate is high and pH is low. This inhibition pre-
vents futile cycling of C0 2 and malate by PEP carboxylase and avoids competition
between PEP carboxylase and Rubisco for C0 2 .

▲ Field of Dreams. These baseball players
were probably studying the biochemistry
of carbon fixation in the corn field.

▲ Cactus is a CAM plant.

472 CHAPTER 15 Photosynthesis

Figure 15.30 ►

Crassulacean acid metabolism (CAM). At

night, C0 2 is taken up, and PEP carboxylase
and NAD®-malate dehydrogenase catalyze
the formation of malate. The phospho-
enolpyruvate required for malate synthesis
is derived from starch. The next day, when
NADPH and ATP are formed by the light
reactions, the decarboxylation of malate in-
creases the cellular concentration of C0 2
that can be fixed by the Calvin cycle. The
decarboxylation of malate occurs by either
of two pathways, depending on the species,
and yields phosphoenolpyruvate, which is
subsequently converted to starch through
gluconeogenesis.

Atmospheric C0 2 -

Carbonic
anhydrase

H 2 0 + C0 2 i=3

Phosphoenol-
pyruvate ~

Glycolysis

HCO , 0 P:

PEP carboxylase

-^Oxaloacetate

Starch

NADH + H®

NAD®-malate

dehydrogenase

NAD @ <-^

NIGHT

DAY

Malate

Jon

Starch

Gluconeo-

genesis

Malate

NAD® — ^

NAD®-malate
dehydrogenase

NADH + H®^

Phosphoenol-

pyruvate

ADP

ATP

AMP + PPj

Pyruvate-

phosphate

dikinase

ATP + P: -

PEP

carboxykinase

CO, —

-Oxaloacetate

> Calvin cycle
NAD(P)H + H® NAD(P)®

Pyruvate

NAD(P)®-malic enzyme

CO 2 > Calvin cycle

Malate

Summary

1. Chlorophyll is the major light- gathering pigment in photosynthe-
sis. When chlorophyll molecules absorb a photon of light, an electron
is promoted to a higher-energy molecular orbital. This elec-
tron can be transferred to an electron transfer chain giving rise to
an electron-deficient chlorophyll molecule.

2. Accessory pigments transfer energy to the special pair of chloro-
phyll molecules by resonance energy transfer.

3. Photosystem II (PSII) complexes contain a type II reaction center.
Electrons are transferred from the special pair of chlorophyll
molecules to a short electron transfer chain consisting of a
chlorophyll, a pheophytin, a bound quinone, and a mobile
quinone.

4. In some bacteria QH 2 molecules from PSII bind to the cytochrome
bc\ complex. Electrons are transferred to cytochrome c and this
process is coupled to the transfer of protons across the membrane
via the Q cycle. Cytochrome c then binds to PSII and transfers
electrons back to the electron-deficient special pair in a cyclic

process of electron transfer. The resulting proton gradient drives
ATP synthesis.

5. Photosystem I (PSI) complexes contain a type I reaction center.
The electron transfer chain consists of two chlorophylls, a phyllo-
quinone, three [Fe-S] clusters, and ferredoxin (or flavodoxin).

6. Reduced ferredoxin is the substrate for ferredoxin: NADP© re-
ductase (FNR), and NADPH is the product of photosystem I
photosynthesis in a noncyclic electron transfer. In some cases,
electrons are passed from ferredoxin to the cytochrome bc\ com-
plex and back to PSI via cytochrome c in a cyclic process of electron
transfer.

7. Cyanobacteria, and chloroplasts, contain coupled photosystems
consisting of PSI, PSII, and cytochrome bf - — a photosynthetic version
of cytochrome bc\. When PSII absorbs a photon of light, electrons
are transferred from PSII to cytochrome bf and plastocyanin.
Plastocyanin resupplies electrons to PSI. When PSI absorbs a
photon of light, excited electrons are used to synthesize NADPH.

Problems 473

In coupled photosystems, PSII is associated with an oxygen evolv-
ing complex (OEC) that catalyzes the oxidation of water to 0 2
and supplies electrons to the PSII special pair.

8. The Z-scheme depicts electron flow during photosynthesis in
terms of the change in reduction potentials of the various compo-
nents of the electron transfer chains.

9. Photosynthesis complexes are concentrated in thylakoid mem-
branes in cyanobacteria. Chloroplasts contain a complex internal
membrane system of thylakoid membranes.

10. The Calvin cycle is responsible for fixing C0 2 into carbohydrates.
The key enzyme is ribulose 1,5-Hsphosphate carboxylase-oxygenase
(Rubisco). Rubisco is an inefficient enzyme that catalyzes car-
boxylation of ribulose 1,5-frzsphosphate. It also catalyzes an
oxygenation reaction.

11. Sucrose and starch are the main products of photosynthetic car-
bohydrate synthesis in plants.

12. Additional carbon-fixation pathways in some plants serve to increase
the concentration of C0 2 at the site of the Calvin cycle reactions.

Problems

1. In plants the transport of a single pair of electrons from P680 to
NADPH is coupled to the accumulation of six protons in the
lumen. This will result in production of 1.5 molecules of ATP
(Section 14.11). Assuming that NADPH ~ 2.5 ATP, this means
that in photosynthesis transport of a pair of electrons through
the complexes produces 1.5 + 2.5 = 4 ATP equivalents. Why
is this process so much more efficient than respiratory electron
transport?

2. The dragonfish is a deepwater species that flashes a red biolumi-
nescent light to illuminate its prey. Although the visual pigments
normally present in the retina of fish are not sensitive enough to
pick up the red light, the dragonfish retina contains other pig-
ments, derived from chlorophyll, that absorb at 667 nm. Suggest
how these chlorophyll pigments might act as a photosensitizer to
aid the dragonfish to detect prey using its own red light beacon,
which other fish cannot see.

3. (a) Ribulose l,5-f7/sphosphate carboxylase-oxygenase (Rubisco)

has been called the “enzyme that feeds the world.” Explain the
basis for this statement.

(b) Rubisco has also been accused of being the world’s most in-
competent enzyme and the most inefficient enzyme in pri-
mary metabolism. Explain the basis for this statement.

4. You frequently see photosynthesis plus the Calvin cycle described as

6 C0 2 + 6 H 2 0 C 6 H 12 O e + 6 0 2

Write a similar equation for the reactions in purple bacteria and
in green sulfur bacteria.

5. (a) Some photosynthesis bacteria use H 2 S as a hydrogen donor

and produce elemental sulfur, whereas others use ethanol and
produce acetaldehyde. Write the net reactions for photosyn-
thesis for these bacteria.

(b) Why is no oxygen produced by these bacteria?

6. Can a suspension of chloroplasts in the dark synthesize glucose
from C0 2 and H 2 0? If not, what must be added for glucose syn-
thesis to occur? Assume that all the components of the Calvin
cycle are present.

7. (a) How many photons are absorbed for every 0 2 molecule pro-

duced in photosynthesis?

(b) How many photons must be absorbed to generate enough
NADPH reducing power for the synthesis of one molecule of
a triose phosphate?

8. The herbicide 3-(3, 4-dichlorophenyl)-l,l-dimethylurea (DCMU)
blocks photosynthetic electron transport from PSII to the cy-
tochrome bf complex.

(a) When DCMU is added to isolated chloroplasts, will both 0 2
evolution and photophosphorylation cease?

(b) If an external electron acceptor that reoxidizes P680* is added,
how will this affect 0 2 production and photophosphorylation?

9. (a) The luminal pH of chloroplasts suspended in a solution of

pH 4.0 reaches pH 4.0 within a few minutes. Explain why
there is a burst of ATP synthesis when the pH of the external
solution is quickly raised to 8.0 and ADP and Pi are added.

(b) If ample ADP and Pi are present, why does ATP synthesis
cease after a few seconds?

10 . Cyclic electron transport may occur simultaneously with non-
cyclic electron transport under certain conditions in chloroplasts.
Is any ATP, 0 2 , or NADPH produced by cyclic electron transport?

11. A plant has been genetically engineered to contain a smaller per-
centage than normal of unsaturated lipids in the thylakoid mem-
branes of the chloroplasts. This genetically changed plant has an
improved tolerance to higher temperatures and also shows im-
proved rates of photosynthesis and growth at 40°C. What major
components of the photosynthesis system might be affected by
changing the lipid composition of the thylakoid membranes?

12 . A compound was added to isolated spinach chloroplasts and the
effect on photosynthetic photophosphorylation, proton uptake,
and noncyclic electron transport determined. Addition of the
compound resulted in an inhibition of photosynthetic pho-
tophosphorylation (ATP synthesis), inhibition of proton uptake,
and an enhancement in noncylic electron transport. Suggest a
mechanism for the compound.

13 . How many molecules of ATP (or ATP equivalents) and NADPH
are required to synthesize (a) one molecule of glucose via photo-
synthetic C0 2 fixation in plants and (b) one glucose residue in-
corporated into starch?

14 . After one complete turn of the Calvin cycle, where will the labeled
carbon atoms from 14 C0 2 appear in (a) glyceraldehyde 3-phos-
phate (b) fructose 6-phosphate, and (c) erythrose 4-phosphate?

15 .

16 .

(a) How many additional ATP equivalents are required to synthesize
glucose from C0 2 in C 4 plants than are required in C 3 plants?

(b) Explain why C 4 plants fix C0 2 much more efficiently than C 3
plants despite the extra ATP needed.

Explain how the following changes in metabolic conditions alter
the Calvin cycle: (a) an increase in stromal pH, and (b) a decrease

in stromal concentration of Mg

,©

474 CHAPTER 15 Photosynthesis

Selected Readings

Pigments

Armstead, I., Donnison, I., Aubry, S., Harper, J.,
Hortensteiner, S., James, C., Mani, J., Moffet, M.,
Ougham, H., Roberts, L., Thomas, A., Weeden, N.,
Thomas, H., and King, I. (2007). Cross-species
identification of Mendel’s I locus. Science 315:73.

Sato Y., Morita R., Nishimura M., Yamaguchi H.,
and Kusaba M. (2007). Mendel’s green cotyledon
gene encodes a positive regulator of the chlorophyll-
degrading pathway. Proc. Natl. Acad. Sci. (USA)
104:14169-14174.

Photosynthetic Electron Transport

Allen, J. R (2004). Cytochrome b^f: structure for
signalling and vectorial metabolism. Trends in
Plant Sci. 9:130-137.

Allen, J. R, and Williams, J. C. (2010). The evolu-
tionary pathway from anoxygenic to oxygenic
photosynthesis examined by comparison of the
properties of photosystem II and bacterial reac-
tion centers. Photosynth. Res. Published online
May 7, 2010: Doi 10. 1007/s 1 1 120-010-9552-x

Amunts, A., Toporik, H., Borovikova, A. B., and
Nelson, N. (2010). Structure determination and
improved model of plant photosystem I. /. Biol.
Chem. 285:3478-3486.

Barber, J., Nield, J., Morris, E. R, and Hankamer, B.
(1999). Subunit positioning in photosystem II
revisited. Trends Biochem. Sci. 24:43-45.

Cramer, W. A., Zhang, H., Yan, J., Kurisu, G., and
Smith, J. L. (2004). Evolution of photosynthesis:
time-independent structure of the cytochrome b 6 f
complex. Biochem. 43:5921-5929.

Cramer, W. A., Zhang, H., Yan, J., Kurisu, G., and
Smith, J. L. (2006). Transmembrane traffic in the
cytochrome b^f complex. Annu. Rev. Biochem.
75:769-790.

Ferreira, K. N., Iverson, T. M., Maghlaoui, K.,
Barber, J., and Iwata, S. (2004). Architecture of the
photosynthetic oxygen-evolving center. Science
303:1831-1838.

Golbeck, J. H. (1992). Structure and function of
photosystem l.Annu. Rev. Plant Physiol. Plant
Mol. Biol. 43:293-324.

Kiihlbrandt, W., Wang, D. N., and Fujiyoshi, Y.
(1994). Atomic model of plant light-harvesting
complex by electron crystallography. Nature 367:
614-621.

Leslie, M. (2009). On the origin of photosynthe-
sis. Science 323:1286-1287.

Muller, M. G., Slavov, C., Luthra, R., Redding,

K. E., and Holzwarth, A. R. (2010). Independent
initiation of primary electron transfer in the
two branches of the photosystem I reaction
center. Proc. Natl. Acad. Sci. (USA)

107:4123-4128.

Nugent, J. H. A. (1996). Oxygenic photosynthesis.
Electron transfer in photosystem I and photosys-
tem II. Eur. J. Biochem. 237:519-531.

Rhee, K.-H., Morris, E. P., Barber, J., and
Kiihlbrandt, W. (1998). Three-dimensional struc-
ture of the plant photosystem II reaction centre at
8 A resolution. Nature 396:283-286.

Staehlin, L. A., and Arntzen, C. J., eds. (1986).
Photosynthesis III: Photosynthetic Membranes
and Light Harvesting Systems. Vol. 19 of

Encyclopedia of Plant Physiology (New York:
Springer- Verlag).

Photophosphorylation

Bennett, J. (1991). Phosphorylation in green plant
chloroplasts. Annu. Rev. Plant Physiol. Plant Mol.
Biol. 42:281-311.

Photosynthetic Carbon Metabolism

Andrews, T. J. and Whitney, S. M. (2003).
Manipulating ribulose frzsphosphate carboxylase/
oxygenase in the chloroplasts of higher plants.
Arch. Biochem. Biophys. 414:159-169.

Bassham, J. A., and Calvin, M. (1957). The Path of
Carbon in Photosynthesis (Englewood Cliffs, NJ:
Prentice Hall).

Edwards, G. E., and Walker, D. (1983). C 3 C 4 :
Mechanisms and Cellular and Environmental Reg-
ulation of Photosynthesis (Berkeley: University of
California Press).

Hartman, F. C., and Harpel, M. R. (1994). Struc-
ture, function, regulation, and assembly of
D-ribulose- 1 ,5- fo'sphosphate carboxylase/oxygenase.
Annu. Rev. Biochem. 63:197-234.

Savage, D. F., Afonso, B., Chen, A. H., and Silver, P. A.
(2010). Spatially ordered dynamics of the bacterial
carbon fixation machinery. Science
327:1258-1261.

Schnarrenberger, C., and Martin, W. (1997). The
Calvin cycle — a historical perspective. Photosynt-
hetica 33:331-345. An outline of the advances
made since the 1950s.

o c

_ o

° o o o

° o

o o

° c

o o

Lipid Metabolism

T he synthesis of lipids is an essential part of cellular metabolism since lipids are
crucial components of cell membranes. In this chapter we describe the path-
ways for synthesis of the major lipids that were described in Chapter 9. The
most important of these pathways is fatty acid synthesis since fatty acids are required in
triacylglycerols. Other important biosynthesis pathways include cholesterol synthesis,
eicosanoid synthesis, and the synthesis of sphingolipids.

Lipids can also be degraded as a normal part of cellular metabolism. The most im-
portant catabolic pathway is that of fatty acid oxidation (/3- oxidation). In this pathway,
long-chain fatty acids are broken down to acetyl CoA. The opposing pathways of fatty
acid biosynthesis and fatty acid oxidation provide another example of how cells handle
energy production and utilization in a manner that’s compatible with the fundamentals
of thermodynamics.

The catabolic pathways of lipid metabolism are part of basic fuel metabolism in an-
imals. Triacylglycerols and glycogen are the two major forms of stored energy. Glycogen
can supply ATP for muscle contraction for only a fraction of an hour. Sustained intense
work, such as the migration of birds or the effort of marathon runners, is fueled by the
metabolism of triacylglycerols. Triacylglycerols are anhydrous and their fatty acids are
more reduced than amino acids or monosaccharides — this makes them very efficient at
storing energy for use later on (Section 9.3). Triacylglycerols are oxidized when the en-
ergy demand increases. In most cases, fat is only used when other energy sources, such
as glucose, are unavailable.

We will begin by examining the fundamental pathways of lipid metabolism — the
ones that are present in all living species. Where necessary, we’ll point out the differ-
ences between the bacterial and the eukaryotic pathways. These differences are minor.
We then go on to describe the absorption and utilization of dietary lipids in mammals,
including the hormonal regulation of lipid metabolism.

16.1 Fatty Acid Synthesis

Fatty acids are synthesized by the repetitive addition of two-carbon units to the growing
end of a hydrocarbon chain. The growing chain is covalently attached to acyl carrier

Derangements of this compli-
cated mechanism of formation
and metabolism of lipids are in
many cases responsible for the
genesis of some of our most im-
portant diseases, especially in the
cardiovascular field. A detailed
knowledge of the mechanisms of
lipid metabolism is necessary to
deal with these medical problems
in a rational manner.

— S. Bergstrom, presentation speech
on awarding the 1964 Nobel Prize in
Physiology or Medicine to Konrad
Bloch and Feodor Lynen

Top: Whereas the polar bear lives off its stored fat for much of the year, the bird uses its fat stores for long flights.

475

476 CHAPTER 16 Lipid Metabolism

Figure 16.1 ►

Outline of fatty acid synthesis.

Initiation stage

(a)

C0 2

Acetyl CoA (C 2 ) — ^ Malonyl CoA (C 3 )

Bacteria !

\ Acetyl ACP (C 2 ) Malonyl ACP (C 3 )
\Eukaryotes

CO?

Acetoacetyl ACP (C 4 )

Elongation stage

(b)

^ » 3-Ketoacyl ACP (C n + 2 )

(C n + 2 ) [Reduction

• CO2 [ Dehydration

Malonyl ACP I Reduction

(C 3 ) ^

Acyl ACP (C n )

c— ch 2 — c
©0 / x o©

Malonate

The regulation of fatty acid metabolism
is described in Section 16.9.

protein (ACP), a protein coenzyme (Section 7.6). The linkage is a thioester as in acetyl
CoA. An overview of fatty acid synthesis is shown in Figure 16.1.

The first steps in the fatty acid synthesis pathway are the production of acetyl ACP
and malonyl ACP from acetyl CoA. (Malonic acid, or malonate, is the name of the stan-
dard C 3 dicarboxylic acid.) The initiation step involves a condensation of acetyl and
malonyl groups to give a four-carbon precursor and C0 2 . This precursor serves as the
primer for fatty acid synthesis. In the elongation stage, the acyl group attached to ACP
(acyl ACP) is extended by two -carbon units donated by malonyl ACP. The product of
the initial condensation (3-ketoacyl ACP) is modified by two reduction reactions and a
dehydration reaction to produce a longer acyl ACP. Acyl ACP then serves as the sub-
strate for additional condensation reactions.

Fatty acid synthesis takes place in the cytosol of all species. In adult mammals it oc-
curs largely in liver cells and adipocytes. Some fatty acid synthesis takes place in special-
ized cells such as mammary glands during lactation.

A. Synthesis of Malonyl ACP and Acetyl ACP

Malonyl ACP is the substrate for fatty acid biosynthesis. It is synthesized in two steps,
the first of which is the carboxylation of acetyl CoA in the cytosol to form malonyl CoA
(Figure 16.2). The carboxylation reaction is catalyzed by the biotin-dependent enzyme
acetyl- CoA carboxylase using a mechanism similar to the reaction catalyzed by pyruvate
carboxylase (Figure 7.20). The ATP-dependent activation of HCOP forms carboxybi-
otin. This reaction is followed by the transfer of activated C0 2 to acetyl CoA, forming
malonyl CoA. These reactions are catalyzed in eukaryotes by a bifunctional enzyme and
the biotin moiety is on a flexible arm that moves between the two active sites. The bac-
terial version of acetyl-CoA carboxylase is a multisubunit enzyme complex containing
biotin carboxylase, biotin carboxylase carrier protein, and a heterodimeric transcar-
boxylase. In all species, acetyl-CoA carboxylase is the key regulatory enzyme of fatty
acid synthesis and the carboxylation reaction is metabolically irreversible.

The second step in the synthesis of malonyl ACP is the transfer of the malonyl moiety
from coenzyme A to ACP. This reaction is catalyzed by malonyl CoA:ACP transacylase
(Figure 16.3). A similar enzyme called acetyl CoA:ACP transacylase converts acetyl CoA to
the acetyl ACP. In most species these are separate enzymes with specificity for malonyl
CoA or acetyl CoA but in mammals the two activities are combined in a bifunctional en-
zyme, malonyl-acetyl transferase (MAT) that’s part of a larger complex (see below).

Figure 16.2 ►

Carboxylation of acetyl CoA to malonyl CoA,
catalyzed by acetyl-CoA carboxylase.

ADP + P:

HCOP + ATP

16.1 Fatty Acid Synthesis 477

B. The Initiation Reaction of Fatty Acid Synthesis

The synthesis of long-chain fatty acids begins with the formation of a four-carbon unit
attached to ACP. This molecule, called acetoacetyl ACP, is formed by condensation of a
two-carbon substrate (acetyl CoA or acetyl ACP) and a three-carbon substrate (malonyl
ACP) with the loss of C0 2 . The reaction is catalyzed by 3-ketoacyl ACP synthase (KAS).

There are several versions of KAS in bacterial cells. One form of the enzyme (KAS
III) is used in the initiation reaction and other versions (KAS I, KAS II) are used in sub-
sequent elongation reactions. Bacterial KAS III uses acetyl CoA for the initial condensa-
tion reaction with malonyl ACP (Figure 16.4).

A two-carbon unit from acetyl CoA is transferred to the enzyme where it is cova-
lently bound via a thioester linkage. The enzyme then catalyzes the transfer of this two-
carbon unit to the end of malonyl ACP creating a four-carbon intermediate and releas-
ing C0 2 . Eukaryotic versions of 3-ketoacyl ACP synthase carry out the same reaction
except that they use acetyl ACP as the initial substrate instead of acetyl CoA.

Recall that synthesis of malonyl CoA involves ATP- dependent carboxylation of acetyl
CoA (Figure 16.2). This strategy of first carboxylating and then decarboxylating a com-
pound results in a favorable free energy change for the process at the expense of ATP con-
sumed in the carboxylation step. A similar strategy is seen in mammalian gluconeogenesis
where pyruvate (C 3 ) is first carboxylated to form oxaloacetate (C 4 ) and then oxaloacetate
is decarboxylated to form the C 3 molecule phosphoenolpyruvate (Section 12.1).

C. The Elongation Reactions of Fatty Acid Synthesis

Acetoacetyl ACP contains the smallest version of a 3-ketoacyl moiety. The “3-keto-” in
the name of this molecule refers to the presence of a keto group at the C-3 position. In
the older terminology this carbon atom was the /3- carbon and the product was called a
/3-ketoacyl moiety. The condensation enzyme is also called /3-ketoacyl ACP synthase.

In order to prepare for subsequent condensation reactions, this oxidized 3-ketoacyl
moiety has to be reduced by the transfer of electrons (and protons) to the C-3 position.
Three separate reactions are required,

o 11

0 OOC— CH 2 — C— S-CoA

Malonyl CoA

Malonyl CoA:ACP HS"ACP
transacylase ^HS"CoA

nJ/

o 11

G ooc— ch 2 — c— s-acp

Malonyl ACP

H 3 C — C — S-CoA
Acetyl CoA

Acetyl CoA:ACP HS"ACP
transacylase ^HS"CoA

\|/

H 3 C — C — S-ACP
Acetyl ACP

▲ Figure 16.3

Synthesis of malonyl ACP from malonyl CoA
and acetyl ACP from acetyl CoA.

Ri — C — CH 2 — R 2

OH H

I I

Rt — C — CH 2 — R 2 > Ri — C = C — R 2

H H

■> Rt — CH 2 — CH 2 — R 2

Reduction Dehydration Reduction (16.1)

The ketone is reduced to the corresponding alcohol in the first reduction. The second step
is the removal of water by a dehydratase producing a C = C double bond. Finally, a

H 3 C — C — CH 2 — C —S-ACP

8 /3 a

4 3 2 1

Acetoacetyl ACP

◄ Figure 16.4

Synthesis of acetoacetyl ACP in bacteria.

478 CHAPTER 16 Lipid Metabolism

Figure 16.5 ►

The elongation stage of fatty acid synthesis.

R represents — CH 3 in acetoacetyl ACP or
[ — CH 2 — CH 2 ] n — CH 3 in other 3-ketoacyl
ACP molecules.

R — C — CH 2 — ( — S-ACP <-
3 Ketoacyl ACP

3-Ketoacyl-ACP

reductase

- NADPH + H®
^ NADP®

R — C — CH 2 — C —S-ACP

D-3-Hydroxyacyl ACP

3-Hydroxyacyl-ACP

dehydratase

H 2 0

R— C= — -S-ACP
H

trans-A 2 -Enoyl ACP

NADPH + H®
NADP®

Enoyl-ACP

reductase

R—CH 2 — CH 2 —C — S-ACP
Acyl-ACP

CO + HS-ACP

3-Ketoacyl-ACP

synthase

'®OOC— CH 2 — C— S-ACP

Malonyl ACP
+ H®

KEY CONCEPT

Malonyl ACP, formed from acetyl CoA, is
the precursor for all fatty acid synthesis.

second reduction adds hydrogens to create the reduced acyl group. This is a common
oxidation-reduction strategy in biochemical pathways. We have seen an example of the
reverse reactions in the citric acid cycle where succinate is oxidized to oxaloacetate
(Figure 13.5).

The specific reactions of the elongation cycle are shown in Figure 16.5. The first re-
duction is catalyzed by 3-ketoacyl ACP reductase (KR). The full name of the dehy-
dratase enzyme is 3-hydroxyacyl ACP dehydratase (DH). The second reduction step is
catalyzed by enoyl ACP reductase (ER). Note that during synthesis the d isomer of the
/3-hydroxy intermediate is formed in an NADPH-dependent reaction. We will see in
Section 16.7 that the l isomer is formed during the degradation of fatty acids.

The final product of the reduction, dehydration, and reduction steps is an acyl ACP
that is two carbons longer. This acyl ACP becomes the substrate for the elongation
forms of 3-ketoacyl ACP synthase (KAS I and KAS II). All species use malonyl ACP as
the carbon donor in the condensation reaction. The elongation reactions are repeated
many times resulting in longer and longer fatty acid chains.

The end products of saturated fatty acid synthesis are 16- and 18-carbon fatty
acids. Larger chain lengths cannot be accommodated in the binding site of the condens-
ing enzyme. The completed fatty acid is released from ACP by the action of a
thioesterase (TE) that catalyzes a cleavage reaction regenerating HS-ACP. For example,
palmitoyl ACP is a substrate for a thioesterase that catalyzes formation of palmitate and
HS-ACP.

H 2 0

Palmitoyl -ACP — - — ->

Thioesterase

Palmitate (C 16 ) + HS-ACP + H 0

(16.2)

16.1 Fatty Acid Synthesis 479

The overall stoichiometry of palmitate synthesis from acetyl CoA and malonyl CoA is

Acetyl CoA + 7 Malonyl CoA + 14 NADPH + 20 H® >

Palmitate + 7 C0 2 + 14 NADP© + 8 HS-CoA + 6 H 2 0

(16.3)

In bacteria, each reaction in fatty acid synthesis is catalyzed by a discrete mono-
functional enzyme. This type of pathway is known as a type II fatty acid synthesis system
(FAS II). In fungi and animals, the various enzymatic activities are localized to individ-
ual domains in a large multifunctional enzyme and the complex is described as a type I
fatty acid synthesis system (FAS I).

The large mammalian polypeptide is about 2500 amino acid residues in length
(Mr = 270 kDa). Fatty acid synthase is a dimer where the two monomers are tightly
bound, creating an enzyme with two sites where the fatty acids are synthesized on either
side of the dimer axis (Figure 16.6). The bottom part of the enzyme in Figure 16.6 con-
tains the condensing activities of malonyl/acetyl transferase (MAT) and 3-ketoacyl ACP
synthase (KAS) that are responsible for adding a new two -carbon unit to the growing
chain. These enzymes attach the fatty acid to a bound ACP phosphopantetheine pros-
thetic group (ACP) that is positioned on a flexible loop. The ACP-bound fatty acid visits
the active sites of the modifying activities: 3-ketoacyl ACP reductase (KR), 3-hydroxyacyl
ACP dehydratase (DH), and enoyl ACP reductase (ER). The fatty acid chain is eventu-
ally released by a thioesterase (TE) activity.

The structures of the ACP domain and the TE domain are not resolved in the crys-
tal structure because they are tethered to the main part of the enzyme by a short stretch
of residues that are intrinsically disordered (Section 4.7D). These flexible domains must
be free to move during the reaction.

D. Activation of Fatty Acids

The thioesterase reaction (Reaction 16.2) results in release of free fatty acids but subse-
quent modifications of these fatty acids require an activation step where they are con-
verted to thioesters of coenzyme A in an ATP-dependent reaction catalyzed by acyl-CoA
synthetase (Figure 16.7). The pyrophosphate released in this reaction is hydrolyzed to
two molecules of phosphate by the action of pyrophosphatase. As a result, two phos-
phoanhydride bonds, or two ATP equivalents, are consumed to form the CoA thioesters
of fatty acids. Bacteria generally have a single acyl-CoA synthetase but in mammals
there are at least four different acyl-CoA synthetase isoforms. Each of the distinct en-
zymes is specific for a particular fatty acid chain length: short (<C 6 ), medium (C 6 to
C 12 ), long (>C 12 ), or very long (>C 16 ). The mechanism of the activation reaction is the
same as that for the synthesis of acetyl CoA from acetate and CoA (Figure 10.13). Activation
of fatty acids is required for their incorporation into membrane lipids (Section 16.2).

E. Fatty Acid Extension and Desaturation

The fatty acid synthase pathway cannot make fatty acids that are longer than 16 or 18
carbons (C 16 or C 18 ). Longer fatty acids are made by extending palmitoyl CoA or
stearoyl CoA in separate extension reactions. The enzymes that catalyze such extensions
are known as elongases and they use malonyl CoA (not malonyl ACP) as the source of
the two-carbon extension unit. An example of an elongase reaction is shown below in
step 2 of Figure 16.8. Long chain fatty acids such as C 2 o and C 22 fatty acids are common
but C 24 and C 26 fatty acids are rare.

Unsaturated fatty acids are synthesized in both bacteria and eukaryotes but the path-
ways are quite different. In type II fatty acid synthesis systems (bacteria) a double bond is
added to the growing chain when it reaches a length of ten carbon atoms. The reaction
is catalyzed by specific enzymes that recognize the C 10 intermediate. For example, 3-
hydroxydecanoyl-ACP dehydratase specifically introduces a double bond at the 2 position
just as in the normal dehydratase reaction during fatty acid synthesis (Figure 16.5). How-
ever, the specific C 10 dehydratase creates a cis- 2-decanoyl ACP and not the trans configu-
ration that serves as a substrate for enoyl ACP reductase.

Axis

▲ Figure 16.6

Mammalian fatty acid synthase. The structure
of the pig {Sus scrofa) enzyme is shown. It
is a large dimer consisting of the following
enzyme activities: malonyl/acetyl transferase
(MAT), 3-ketoacyl ACP synthase (KAS),
3-ketoacyl ACP reductase (KR), 3-hydroxya-
cyl ACP dehydratase (DH), enoyl ACP reduc-
tase (ER), and thioesterase (TE). The fatty
acid chain is attached to a bound ACP co-
factor (ACP). The structures of the ACP and
TE domains are not resolved because they
are bound to a flexible tether. [PDB 2VZ9]

R— COO G + HS-CoA
Fatty acid
^ATP

Acyl-CoA synthetase

AaMP + PPi

R — C— S-CoA
Acyl CoA

▲ Figure 16.7
Activation of fatty acids.

480 CHAPTER 16 Lipid Metabolism

Figure 16.8 ►

Elongation and desaturation reactions in the
conversion of linolenoyl CoA to arachidonoyl
CoA.

The nomenclature of unsaturated fatty
acids is described in Section 9.2.

Linolenoyl CoA (18:2 c/s, c/'s-A 9, 1 2 )

12 9 6

y-Linolenoyl CoA (18:3 all c/'s-A 6,9,12 )

G 11

u ooc — ch 2 — c —

Malonyl CoA
► C0 2 + HS-CoA

S-CoA

Reduction, ^-2 NADH + 2H©
dehydration

reduction ^ 2 NAD © + H 2 Q

Eicosatrienoyl CoA (20:3 all c/'s-A 8,11,14 )

14 11 8 5

S-CoA

Arachidonoyl CoA
(20:4 all c/s- A 5,8,11,14 )

Subsequent elongation of this unsaturated fatty acid proceeds by the normal fatty
acid synthase pathway except that a specific 3-ketoacyl-ACP synthase enzyme recog-
nizes the unsaturated fatty acid in the condensation reaction. The final products will be
16:1 A 8 and 18:1 A 10 unsaturated fatty acids. These products can be further modified to
create polyunsaturated fatty acids (PUFAs) in bacteria. The chains can be extended by
elongase enzymes and additional double bonds are introduced by a class of enzymes
called desaturases. Bacteria contain a huge variety of PUFAs that serve to increase the
fluidity of membranes when species encounter low temperatures (Section 9.9). For ex-
ample, many species of marine bacteria synthesize 20:5 and 22:6 PUFAs. Up to 25% of
the membrane fatty acids are large polyunsaturated fatty acids in these species.

The introduction of a double bond during synthesis of fatty acids is not possible in
eukaryotes since they employ a type I fatty acid synthase. This fatty acid synthase con-
tains a single 3-ketoacyl-ACP synthase (KAS) activity that is part of a large multifunc-
tional protein. The eukaryotic KAS active site does not recognize unsaturated fatty acid
intermediates and could not extend them if they were created at the C 10 step as in bacteria.

16.2 Synthesis of Triacylglycerols and G lycerophosphol i pids 481

Consequently, eukaryotes synthesize unsaturated fatty acids entirely by using desat-
urases that act on the completed fatty acid derivatives palmitoyl CoA and stearoyl CoA.

Most eukaryotic cells contain various desaturases that catalyze the formation of
double bonds as far as 15 carbons away from the carboxyl end of a fatty acid. For exam-
ple, palmitoyl CoA is oxidized to its 16:1 A 9 analog that can be hydrolyzed to form the
common fatty acid palmitoleate. Polyunsaturated fatty acids are synthesized by the se-
quential action of different, highly specific desaturases. In most cases, the double bonds
are spaced at 3 -carbon intervals as in synthesis of a-linolenate in plants.

18:0 (stearoyl CoA) > 18:1A 9 > 18:2A 9,12 (linolenoyl CoA) >

Q i i c- (1 6.4)

18:3A 9,12,15 (u-linolenoyl CoA)

Mammalian cells do not contain a desaturase that acts beyond the C-9 position and
they are not able to synthesize linoleate or ct-linolenate. However, PUFAs with double
bonds at the 12 position are absolutely essential for survival since they are precursors
for synthesis of important eicosanoids such as prostaglandins. Because they lack a A 12
desaturase, mammals must obtain linoleate from the diet. This is an essential fatty acid
in the human diet. Deficiencies of a-lineolate are rare since most food contains ade-
quate quantities. Plants, for example, are rich sources of PUFAs. Nevertheless, the com-
position of many “vitamin” supplements will include linoleic acid.

Mammals can convert dietary linoleate (activated to linolenoyl CoA) to arachi-
donoyl CoA (20:4) by a series of desaturation and elongation reactions (Figure 16.8).
(Arachidonate derived from phospholipids is a precursor of eicosanoids, Section 16.3.)
This pathway illustrates typical examples of elongase and desaturase activity in the syn-
thesis of complex PUFAs. The intermediate y-linolenoyl CoA (18:3) in the arachidonate
pathway can undergo elongation and desaturation to produce C 20 and C 22 polyunsatu-
rated fatty acids. Note that the double bonds of polyunsaturated fatty acids are not con-
jugated but are interrupted by a methylene group. Thus, a A 9 double bond, for example,
directs insertion of the next double bond to the A 6 position or the A 12 position.

16.2 Synthesis of Triacylglycerols and
Glycerophospholipids

Most fatty acids are found in esterified forms as triacylglycerols or glycerophospholipids
(Sections 9.3 and 9.4). Phosphatidate is an intermediate in the synthesis of triacylglyc-
erols and glycerophospholipids. It is formed by transferring the acyl groups from fatty
acid CoA molecules to the C-l and C-2 positions of glycerol 3-phosphate (Figure 16.9).
Glycerol 3 -phosphate is synthesized from dihydroxyacetone phosphate in a reduction
reaction catalyzed by glycerol 3 -phosphate dehydrogenase. We encountered this enzyme
when we discussed NADH shuttle mechanisms in Chapter 14 (Section 14.12).

The lipid synthesis reactions are catalyzed by two separate acyltransferases that use
fatty acyl CoA molecules as the acyl group donors. The first acyltransferase is glycerol-3 -
phosphate acyltransferase. It catalyzes esterification at C-l of glycerol 3 -phosphate to
form 1-acylglycerol 3 -phosphate (lysophosphatidate) and it exhibits a preference for satu-
rated fatty acyl chains. The second acyltransferase is 1-acylglycerol- 3 -phosphate acyltrans-
ferase and it catalyzes esterification at C-2 of 1-acylglycerol 3 -phosphate. This enzyme

▲ Linoleate. Linoleate is an essential
component of the human diet.

In addition to the essential fatty acids,
mammalian diets must supply a number
of essential vitamins (Chapter 7) and
essential amino acids (Chapter 17).

In the older biochemistry literature tri-
acylglycerols were called triglycerides
(Section 9.3).

'CH,— OH Ri — C— S-CoA HS-CoA

O O

II ii

CH 2 — O— C — R, R 2 —C— S-CoA HS-CoA

CH,

-c-

HO— CH

3 CH 2 — uru 3 -
Glycerol 3-phosphate

-OPO,®

Glycerol-3-phosphate

acyltransferase

HO — CH 2

CH 2 — 0P0 3 ®

1 -Acylglycerol-3-phosphate
acyltransferase

FG — C — O — CH

1-Acylglycerol 3-phosphate
(Lysophosphatidate)

CH 2 — 0P0 3 ®
Phosphatidate

▲ Figure 16.9

Formation of phosphatidate. Glycerol 3-phosphate acyltransferase catalyzes esterification at C-l of glycerol 3-phosphate. It has a preference for
saturated acyl chains. l-Acylglycerol-3-phosphate acyltransferase catalyzes esterification at C-2 and has a preference for unsaturated acyl chains.

482 CHAPTER 16 Lipid Metabolism

O CH 2 — o— c — R,

II I

R 2 — C— O— CH

CH 2 — OP0 3 ®
Phosphatidate

Phosphatidate

phosphatase ^ p

-h 2 o

CH 2 — o— C — Rt

R 3 — C— S-CoA
CoA-SH <

R 2 — C — O— CH

Diacylglycerol

acyltransferase

£l_l q |_| Phosphoethanolamine

2 transferase

1,2-Diacylglycerol

CDP — O — (CH 2 ) 2 — NH 3

CDP-ethanolamine

-> CMP

CDP — O — (CH 2 ) 2 — N(CH 3 ) 3 -

O CDP-choline

O CH 2 — O — C — Ri

II I

R 2 — c — o — CH

ch 2 — o— c — r 3

Triacylglycerol

Phosphocholine

transferase

CMP^-

o ch 2 — o— c — r,

II I

R 2 — c — o— CH O

II 0

ch 2 — o— p — o— ch 2 — ch 2 — nh 3

0 (

I©

O M ethylations / Phosphatidylethanolamine

CH 2 — O — C — Ri

R ? — C — O — CH

ch 2 — o— p — o— ch 2 — ch 2 — N(CH 3 ) 3

',©

Phosphatidylcholine

▲ Figure 16.10

Synthesis of triacylglycerols and neutral phospholipids. The formation of triacylglycerols, phosphatidylcholine, and phosphatidylethanolamine proceeds
via a diacylglycerol intermediate. A cytosine-nucleotide derivative donates the polar head groups of the phospholipids. Three enzymatic methylation
reactions, in which S-adenosylmethionine is the methyl-group donor, convert phosphatidylethanolamine to phosphatidylcholine.

prefers unsaturated chains. The product of the two reactions is a phosphatidate, one of a
family of molecules whose specific properties depend on the attached acyl groups.

The formation of triacylglycerols and neutral phospholipids from phosphatidate be-
gins with a dephosphorylation catalyzed by phosphatidate phosphatase (Figure 16.10).
The product of this reaction is a 1,2 -diacylglycerol that can be directly acylated to form a
triacylglycerol. Alternatively, 1,2 -diacylglycerol can react with a nucleotide-alcohol de-
rivative, such as CDP-choline or CDP-ethanolamine (Section 7.3), to form phos-
phatidylcholine or phosphatidylethanolamine, respectively. These derivatives are formed
from CTP by the general reaction

CTP + Alcohol phosphate > CDP-alcohol + PPj (16.5)

Phosphatidylcholine can also be synthesized by methylation of phosphatidyl-
ethanolamine by S-adenosylmethionine (Section 7.3).

Phosphatidate is also the precursor of acidic phospholipids. In this pathway, phos-
phatidate is first activated by reacting with CTP to form CDP-diacylglycerol with the re-
lease of pyrophosphate (Figure 16.1 1). In some bacteria, the displacement of CMP by ser-
ine produces phosphatidylserine. In both prokaryotes and eukaryotes, displacement of
CMP by inositol produces phosphatidylinositol. Phosphatidylinositol can be converted to
phosphatidylinositol 4-phosphate (PIP) and phosphatidylinositol 4,5-frisphosphate

16.3 Synthesis of Eicosanoids 483

Figure 16.11 ►

Synthesis of acidic phospholipids.

Phosphatidate accepts a cytidylyl group
from CTP to form CDP— diacylglycerol. CMP
is then displaced by an alcohol group of
serine or inositol to form phosphatidylserine
or phosphatidylinositol, respectively.

O CH 2 — o — c — Rt
R 2 — c— o — CH O

I II Q

ch 2 — o— p —

Phosphatidate

CTP:phosphatidate

cytidylyltransferase

CTP

PPi

O CH 2 — o — c — FT

II I

R 2 — c — o — CH O

I II

ch 2 — o — p— o— ch 2 — CH — COO^

o°

Phosphatidylserine

O CH 2 — O — C — Ri

II I

R 2 — c — o — CH O

Phosphatidylinositol

(PIP 2 ) through successive ATP-dependent phosphorylation reactions. Recall that PIP 2 is
the precursor of the second messengers inositol 1,4,5-tnsphosphate (IP 3 ) and diacylglyc-
erol (Section 9.1 ID).

Most eukaryotes use a different pathway for the synthesis of phosphatidylserine. It
is formed from phosphatidylethanolamine via the reversible displacement of ethanolamine
by serine, catalyzed by phosphatidylethanolamine:serine transferase (Figure 16.12).
Phosphatidylserine can be converted back to phosphatidylethanolamine in a decar-
boxylation reaction catalyzed by phosphatidylserine decarboxylase.

16.3 Synthesis of Eicosanoids

There are two general classes of eicosanoids: prostaglandins + thromboxanes, and
leukotrienes. Arachidonate (20:4 A 5,8,11,14 ) is the precursor of many eicosanoids. Recall
that arachidonate is synthesized from linoleoyl CoA (18:2 A 9,12 ) in a pathway that re-
quires a A 6 desaturase, an elongase, and a A 5 desaturase as shown in Figure 16.8.

Prostaglandins are synthesized by the cyclization of arachidonate in a reaction cat-
alyzed by a bifunctional enzyme called prostaglandin endoperoxide H synthase (PGHS).

484 CHAPTER 16 Lipid Metabolism

Figure 16.12 ►

Interconversions of phosphatidylethanolamine
and phosphatidylserine.

O CH 2 — O — C — Rt

II I

R 2 — c— o — CH O

ch 2 — o— p — o— ch 2 — ch 2 — nh 3

0 (

Phosphatidylethanolamine

HO — CH 2 — CH — COO°

Serine

Phosphatidylethanolamine:serine

transferase

Ethanolamine
HO — CH 2 — CH 2 — NH 3 ©

co 2

Phosphatidylserine decarboxylase

O CH 2 — O — C — R,

II I

R 2 — C — O— CH O

©NH,

CH 2 — O— P— O — CH 2 — CH — COO°

o 0

Phosphatidylserine

The enzyme is bound to the inner surface of the endoplasmic reticulum through a
cluster of hydrophobic a helices that penetrate one of the lipid bilayers (Figure 16.13).
The cyclooxygenase (COX) activity of the enzyme catalyzes the formation of a hydroper-
oxide (prostaglandin G 2 ). The PGHS enzyme contains a second active site for a hy-
droperoxidase activity that rapidly converts the unstable hydroperoxide to prostaglandin
H 2 (Figure 16.14). This product is converted to various short-lived regulatory molecules
including prostacyclin, prostaglandins, and thromboxane A 2 . Unlike hormones, which are

BOX 16.1 sn-GLYCEROL 3-PHOSPHATE

One of the precursors for synthesis of triacylglycerols is glyc-
erol 3-phosphate shown as a Fischer projection in Figure 16.9.
This molecule could also be accurately drawn upside down as
glycerol 1 -phosphate. This changes the stereochemical nam-
ing convention from L to D. Similarly, D-glycerol 3 -phosphate
and L-glycerol 1 -phosphate are different names for the same
molecule.

Having different names for the same molecule could
lead to confusion since the glycerol phosphate precursor is a
prochiral molecule meaning that modified lipids will have
different stereochemical names depending on whether you
start with L-glycerol 3-phosphate or D-glycerol 1 -phosphate.
In order to avoid this, a new convention is introduced to
number the carbon atoms. In a Fischer projection where the
hydroxyl group on C-2 is on the left, the “top” carbon atom
becomes C-l and the “bottom” one is C-3. Thus, L-glycerol
3-phosphate becomes sn-glycerol 3-phosphate where “sn”
stands for Stereochemical numbering.

The accurate name for the triglyceride precursor is
sn-glycerol 3 -phosphate in most cases. In archaebacteria the
precursor is sn - glycerol 1 -phosphate (Box 9.5).

ch 2 oh

ch 2 opo 3 h 2

= — OH

ch 2 opo 3 h 2

CH 2 OH

L-Glycerol 3-phosphate

D-Glycerol 1 -phosphate

sn-Glycerol 3-phosphate

CH 2 OH

ch 2 opo 3 h 2

— OH

= — H

ch 2 opo 3 h 2

CH 2 OH

D-Glycerol 3-phosphate

L-Glycerol 1 -phosphate

sn-Glycerol 1 -phosphate

16.3 Synthesis of Eicosanoids 485

Cytoplasm

◄ Figure 16.13

Prostaglandin endoperoxide H synthase (PGHS,
COX-1). This enzyme is a dimer bound to the
inner membrane of the endoplasmic reticu-
lum. The active sites of the cyclooxygenase
and hydroperoxidase activities are located in
the large cleft at the bottom of the enzyme,
[from sheep, Ovis aries, PDB 1PRH]

Arachidonate

Prostaglandin H synthase:
cyclooxygenase activity

2 0 2

[inhibited by aspirin]

Prostaglandin G 2

00 H

5-Hydroperoxy-A 6,8,11,14 -eicosatetraenoate

Prostaglandin endoperoxide H synthase:
hydroperoxidase activity

Dehydrase

^ h 2 o

coo°

Prostacyclin

Other prostaglandins

Thromboxane A 2

▲ Figure 16.14

Major pathways for the formation of eicosanoids. The prostaglandin H synthase (PGHS) pathway leads to prostaglandin H 2 that can be converted to
prostacyclin, thromboxane A 2 and a variety of prostaglandins. The lipoxygenase pathway shown produces leukotriene A 4 a precursor of some other
leukotrienes. The cyclooxygenase activity of PGHS is inhibited by aspirin.

486 CHAPTER 16 Lipid Metabolism

▲ The bark of willow trees is a natural source
of salicylates.

produced by glands and travel in the blood to their sites of action, eicosanoids typically
act in the immediate neighborhood of the cell in which they are produced. For example,
thromboxane A 2 is produced by blood platelets and it leads to platelet aggregation and
blood clots and constriction of the smooth muscles in arterial walls causing localized
changes in blood flow. The uterus produces contraction- triggering prostaglandins dur-
ing labor. Eicosanoids also mediate pain sensitivity, inflammation, and swelling.

Recall that linoleate must be supplied in the human diet, usually from plants, in
order to support the synthesis of arachidonate and eicosanoids. One of the reasons why
linoleate is essential is because it’s required for synthesis of prostaglandins and
prostaglandins are necessary for survival.

Aspirin blocks production of some eicosanoids and thus relieves the symptoms of
pain and reduces fever. The active ingredient of aspirin, acetylsalicylic acid, irreversibly
inhibits COX activity by transferring an acetyl group to an active-site serine residue of
the bifunctional enzyme. By blocking the activity of COX, aspirin prevents the forma-
tion of a variety of eicosanoids that are synthesized after the COX reaction. Aspirin was
first developed as a marketable drug in 1897 but other salicylates have long been used in
the treatment of pain. The ancient Greeks, for example, used the bark of willow trees for
pain relief. Willow bark is a natural source of salicylates.

The second class of eicosanoids are the products of reactions catalyzed by lipoxyge-
nases. In Figure 16.14, arachidonate lipoxygenase is shown catalyzing the first step in
the pathway leading to leukotriene A 4 . (The term triene refers to the presence of three
conjugated double bonds.) Further reactions produce other leukotrienes, such as the
compounds once called the “slow-reacting substances of anaphylaxis” (allergic re-
sponse) that are responsible for the occasionally fatal effects of exposure to antigens.

BOX 16.2 THE SEARCH FOR A REPLACEMENT FOR ASPIRIN

Most natural salicylates have serious side effects. They cause
inflamation of the mouth, throat, and stomach and they taste
horrible. Aspirin avoids most of these side effects, which is
why it became such a popular drug when it was first intro-
duced. However, aspirin can cause dizziness, ringing in the
ears, and bleeding or ulcers of the stomach lining. There are
two different forms of PGHS (also called COX after their cy-
clooxygenase activity). COX-1 is a constitutive enzyme that
regulates secretion of mucin in the stomach, thus protecting
the gastric wall. COX-2 is an inducible enzyme that promotes
inflammation, pain, and fever. Aspirin inhibits both isozymes.

There are many other nonsteroidal anti-inflammatory
drugs (NSAIDS) that inhibit COX activity. Aspirin is the only
one that inhibits by covalent modification of the enzyme. The
others act by competing with arachidonate for binding to the

COX active site. Ibuprofen (Advil®), for example, binds rap-
idly, but weakly, to the active site and its inhibition is readily
reversed when drug levels drop. Acetaminophen (Tylenol®) is
an effective inhibitor of COX activity in intact cells.

Physicians would like to have a drug that selectively in-
hibits COX-2 and not COX-1. Such a compound would not
cause stomach irritation. A number of specific COX-2 in-
hibitors have been synthesized and many are currently avail-
able for patients. These drugs, while expensive, are important
for patients with arthritis who must take pain killers on a regu-
lar basis. In some cases, the new NSAIDS have been associated
with increased risk of cardiovascular disease and they have
been taken off the market (e.g., Vioxx®). X-ray crystallographic
studies of COX-2 and its interaction with these inhibitors has
aided the search for even better replacements for aspirin.

ch 3 x ch 3

Ibuprofen

Rofecoxib (Vioxx®)
(COX-2 specific NSAID)

16.4 Synthesis of Ether Lipids 487

16.4 Synthesis of Ether Lipids

Ether lipids have an ether linkage in place of one of the usual ester linkages (Section 9.4).
The pathway for the formation of ether lipids in mammals begins with dihydroxyacetone
phosphate (Figure 16.15). First, an acyl group from fatty acyl CoA is esterified to the oxy-
gen atom at C-l of dihydroxyacetone phosphate producing 1-acyldihydroxyacetone
phosphate. Next, a fatty alcohol displaces the fatty acid to produce 1-alkyldihydroxyace-
tone phosphate. The keto group of this compound is then reduced by NADPH to form
l-alkylglycero-3-phosphate. This reduction is followed by esterification at C-2 of
the glycerol residue to produce l-alkyl-2-acylglycero-3 -phosphate. The subsequent
reactions — dephosphorylation and addition of a polar head group (either choline or
ethanolamine) — are the same as those shown earlier in Figure 16.10. Plasmalogens,
which contain a vinyl ether linkage at C-l of the glycerol backbone (Figure 9.9), are
formed from alkyl ethers by oxidation of the alkyl ether linkage. This reaction is cat-
alyzed by an oxidase that requires NADH and 0 2 . The oxidase is similar to the acyl-CoA
desaturases (Figure 16.8) that introduce double bonds into fatty acids.

In eukaryotes, ether lipids are not as common as the glycerophospholipids containing
ester linkages although some species and some tissues have membranes that are enriched
in plasmalogens. Ether lipids are more common in bacteria, especially in archaebacteria
where the majority of membrane lipids are ether lipids (Box 9.5).

CH 2 — OH
C=0

ch 2 — opo 3 ©

Dihydroxyacetone

phosphate

Dihydroxyacetone

phosphate

acyltransferase

R — C— S-CoA
^HS-CoA

O CH 2 — O — CH 2 — CH^

R 2 — C — O— CH O

I II

ch 2 — o— p — o— ch 2 ch 2

O©

1-Alkyl-2-acylglycero-3-phosphocholine

CMP

Phosphocholine transferase
CDP-choline

CH 2 — O — C — R
C=0

ch 2 — opo 3 ©

1-Acyldihydroxyacetone

phosphate

O CH 2 — O — CH 2 CH 2 R<|

R 2 — C — O— CH

CH 2 — OH

1 -Alkyl-2-acylglycerol

1-Alkyl-

dihydroxyacetone

phosphate

synthase

■ HO — CH 2 CH 2 R.|

V > 0 O — C — R + H©

Phosphatase

H 2 0

ch 2 — o— ch 2 ch 2 r 1
c = o

CH 2 — 0P0 3 ©
1-Alkyldihydroxyacetone
phosphate

O CH 2 — O — CH 2 CH 2 R!

R 2 — C — O— CH

CH 2 — OPO 3 ©

1-Alkyl-2-acylglycero-3-phosphate

1-Alkyl-
dihydroxyacetone

phosphate L NADp ©
oxidoreductase

NADPH + H 0

ch 2 — o — CH 2 CH 2 R-!
HO — CH

CH 2 — 0P0 3 ®

HS-CoA

1-Alkylglycerophosphate

acyltransferase

R 2 —C — S-CoA

1-Alkylglycero-3-phosphate

Figure 16.15 ▼

Synthesis of ether lipids. Plasmalogens
are synthesized from ether lipids by
the formation of a double bond at
the position marked with a red arrow.

,N(CH 3 ) 3

488

CHAPTER 16 Lipid Metabolism

16.5 Synthesis of Sphingolipids

Sphingolipids are membrane lipids that have sphingosine (a C 18 unsaturated amino al-
cohol) as their structural backbone (Figure 9.10). In the first step of sphingolipid
biosynthesis, serine (a C 3 unit) condenses with palmitoyl Co A, producing 3-ketosphin-
ganine and C0 2 (Figure 16.16). Reduction of 3-ketosphinganine by NADPH produces
sphinganine. Next, a fatty acyl group is transferred from acyl CoA to the amino group
of sphinganine in an N-acylation reaction. The product of this reaction is dihydroce-
ramide, or ceramide without the characteristic double bond between C-4 and C-5 of a
typical sphingosine. This double bond is introduced in a reaction catalyzed by dihydro-
ceramide A 4 -desaturase, an enzyme that is similar to other desaturases that we have en-
countered. The final product is ceramide (N-acylsphingosine).

Ceramide is the source of all the other sphingolipids. It can react with phosphatidyl-
choline to form sphingomyelin or with a UDP-sugar to form a cerebroside. Complex
sugar-lipid conjugates, gangliosides, can be formed by reaction of a cerebroside with ad-
ditional UDP-sugars and CMP-N-acetylneuraminic acid (Figure 9.12). Gangliosides are
found in the outer leaflet of the plasma membrane, as are most glycolipids.

16.6 Synthesis of Cholesterol

The steroid cholesterol is an important component of many membranes (Section 9.8)
and a precursor of steroid hormones and bile salts in mammals. All the carbon atoms in
cholesterol come from acetyl CoA, a fact that emerged from early radioisotopic labeling
experiments. Squalene, a C 30 linear hydrocarbon, is an intermediate in the biosynthesis
of the 27-carbon cholesterol molecule. Squalene is formed from 5-carbon units related
to isoprene. The stages in the cholesterol biosynthesis pathway are

Acetate (C 2 ) » Isoprenoid (C 5 ) > Squalene (C 30 ) > Cholesterol (C 27 ) (16.6)

A. Stage 1: Acetyl CoA to Isopentenyl Diphosphate

The first step in cholesterol synthesis is sequential condensation of three molecules of
acetyl CoA. These condensation steps are catalyzed by acetoacetyl-CoA thiolase and
HMG-CoA synthase. The product, HMG CoA, is then reduced to mevalonate in a reac-
tion catalyzed by HMG-CoA reductase (Figure 16.17). This is the first committed step in
cholesterol synthesis. Mevalonate is converted to the C 5 compound isopentenyl diphos-
phate by two phosphorylations followed by decarboxylation. The conversion of three
molecules of acetyl CoA to isopentenyl diphosphate requires energy in the form of three
ATP and two NADPH. In addition to its role in cholesterol synthesis, isopentenyl diphos-
phate is an important donor of isoprenyl units for many other biosynthesis reactions.

Many species of bacteria have a completely different, mevalonate- independent path-
way for synthesis of isopentyl diphosphate. The initial precursors in this pathway are glyc-
eraldehyde 3-phosphate + pyruvate and not acetyl CoA. The mevalonate-independent
pathway is more ancient than the mevalonate- dependent pathway shown here.

B. Stage 2: Isopentenyl Diphosphate to Squalene

Isopentenyl diphosphate is converted to dimethylallyl diphosphate by a specific iso-
merase called isopentenyl diphosphate isomerase (IDI). The two isomers are then joined
in a head-to-tail condensation reaction catalyzed by prenyl transferase (Figure 16.18).
The products of this reaction are a C 10 molecule (geranyl diphosphate) and pyrophos-
phate. A second condensation reaction, also catalyzed by prenyl transferease, produces
the important C 15 intermediate, farnesyl diphosphate. The condensation of isoprenyl
units produces a characteristic branched hydrocarbon with regularly spaced double
bonds at the branch position. These isoprene units (Figure 9.13) are present in a number
of important cofactors.

Two molecules of farnesyl diphosphate are joined in a head-to-head condensation
reaction to form the C 30 molecule squalene. Pyrophosphate, whose hydrolysis drives
reaction equilibria toward completion, is produced in three steps in the squalene synthesis
pathway. Note that all double bonds in squalene are trans.

Mitochondrial isozymes of acetoacetyl-
CoA thiolase and HMG-CoA synthase
are involved in the synthesis of ketone
bodies (Section 16.1 1).

KEY CONCEPT

Isopentenyl diphosphate is the precursor
for synthesis of all isoprenoids.

16.6 Synthesis of Cholesterol 489

coo°

H 3 N — CH O

I II

CH 2 OH CoA-S — c — (ch 2 ) 14 — ch 3

Serine Palmitoyl CoA

C-(CH 2 ) 14 -CH 3

H 3 N — CH

CH 2 OH

NADPH+H© NADP©

3-Ketosphinganine

reductase

3-Ketosphinganine

◄ Figure 16.16
Synthesis of sphingolipids.

CH-(CH 2 ) 14 -CH 3

H 3 N — CH

ch 2 oh

Sphinganine

/ — R 1 — C — S-CoA

Sphinganine

A/-acyltransferase

"^HS-CoA

R i

OH R

I H

O CH — C = C — (CH 2 ) 12 — CH 3

II I H

— C — N — CH o

H I II ©

CH 2 — O — P — O— CH 2 CH 2 N(CH 3 ) 3

o©

r .. Phosphatidylcholine

Sphingomyelin /

1,2-Diacylglycerol

O CH — (CH 2 ) 14 — CH 3

II I

! — C — N — CH

H |
ch 2 oh

A/-Acylsphinganine

(Dihydroceramide)

0 2

h 2 o

NADH + H©

Dihydroceramide

A 4 -desaturase

>NAD©

I H

O CH — C=C— (CH 2 ) 12 — CH

II I H

— C — N — CH

H I

ch 2 oh

Ceramide

Cerebroside

(Galactocerebroside)

490 CHAPTER 16 Lipid Metabolism

h 3 c— c— S-C oA h 3 c— c— S-CoA
Acetyl CoA Acetyl CoA

\ ^ , /

Acetoacetyl CoA
thiolase

H,C

CoA-S — C — CH 3
Acetyl CoA

H 3 C — C—CH 2 —C — S-CoA
Acetoacetyl CoA

H,C

C —

ch 2 — ch 2 — o— p— o— p— 0°

6°

Isopentenyl diphosphate

h 2 o

H MG -CoA synthase

H + + HS-CoA

©,

OOC — CH 2 — C — CH 2 — C— S-CoA
CH 3

3-Hydroxy-3-methylglutaryl CoA (HMG CoA)
- 2NADPH + 2H @

ADP + Pj
ATP

HCO

Mevalonate-5-diphosphate

decarboxylase

©,

ooc— ch 2 — c— ch 2 — ch 2 — o— p— o— p— 0 °

I©

HMG-CoA

reductase

ch 3 o'

Mevalonate-5-diphosphate

\©

©,

OOC — CH,

2NADP®
^ HS-CoA
OH

c — ch 2 -
ch 3

Mevalonate

CHo — OH

ATP ADP

Mevalonate

kinase

ADP <
ATP -
OH

Phosphomevalonate

kinase

©,

ooc—

ch 2 — c — ch 2 — ch 2 — o— p— o 0

ch 3

Mevalonate-5-phosphate

▲ Figure 16.17

Stage I of cholesterol synthesis: formation of isopentenyl diphosphate. The condensation of three
acetyl CoA molecules leads to HMG CoA, which is reduced to mevalonate. Mevalonate is then con-
verted to the five-carbon molecule isopentenyl diphosphate via two phosphorylations and one
decarboxylation.

C. Stage 3: Squalene to Cholesterol

The steps between squalene and the first fully cyclized intermediate, lanosterol, include
the addition of a hydroxyl group followed by a concerted series of cyclizations to form
the four-ring steroid nucleus (Figure 16.19). Lanosterol accumulates in appreciable
quantities in cells that are actively synthesizing cholesterol. The conversion of lanosterol
to cholesterol occurs via two pathways, both involving many steps.

D. Other Products of Isoprenoid Metabolism

A multitude of isoprenoids are synthesized from cholesterol or its precursors. Isopen-
tenyl diphosphate, the C 5 precursor of squalene, is the precursor of a large number of
other products, such as quinones; the lipid vitamins A, E, and K; carotenoids; terpenes;
the side chains of some cytochrome heme groups; and the phytol side chain of chlorophyll
(Figure 16.20). Many of these isoprenoids are made in bacteria, which do not synthesize

◄ Konrad Bloch (1912-2000) (top) and Feodor Lynen (1911-1979) (bottom) received the Nobel Prize
in Physiology or Medicine in 1964 “for their discoveries concerning the mechanism and regulation
of the cholesterol and fatty acid metabolism”.

16.6 Synthesis of Cholesterol 491

H,C.

C — CH,— CH ? — O — P — O— P — 0 (

H,C

Isopentenyl diphosphate

Isopentenyl

diphosphate

isomerase

h 3 c

o o

\ p. II II ©

C = CH — CH,— O— P — O — P — 0°

,©

Dimethylallyl diphosphate

PP,^

H 3 C

©CH,

h 3 c

\=ch

h 3 c / ch 2 -op 2 o 6

N.C-r-C — H

V Va

h 2 c h

Isopentenyl diphosphate

Prenyl transferase

H©^A

OP,O fi ©

Isopentenyl

diphosphate

(C 5 )

Prenyl transferase

PPr

◄ Figure 16.18

Condensation reactions in the second stage of
cholesterol synthesis.

▲ Figure 16.19

Final stage of cholesterol synthesis: squalene
to cholesterol. The conversion of lanosterol
to cholesterol requires up to 20 steps.

Squalene synthase

NADPH + H©
NADP©

Squalene

(C30)

492 CHAPTER 16 Lipid Metabolism

BOX 16.3 REGULATING CHOLESTEROL LEVELS

The HMG-CoA reductase reaction appears to be the princi-
pal site for the regulation of cholesterol synthesis. HMG-CoA
reductase has three regulatory mechanisms — covalent modi-
fication, repression of transcription, and control of degrada-
tion. Short-term control is effected by covalent modification:
HMG-CoA reductase is an interconvertible enzyme that is
inactivated by phosphorylation. This phosphorylation is cat-
alyzed by an unusual AMP-activated protein kinase that can
also catalyze the phosphorylation and concomitant inactiva-
tion of acetyl-CoA carboxylase (Section 16.9). The action of
the kinase appears to decrease the ATP-consuming synthesis
of both cholesterol and fatty acids when AMP levels rise. The
amount of HMG-CoA reductase in cells is also closely regu-
lated. Cholesterol (endogenous cholesterol delivered by
plasma lipoproteins or dietary cholesterol delivered by chy-
lomicrons) can repress transcription of the gene that encodes

HMG-CoA reductase. In addition, high levels of cholesterol
and its derivatives increase the rate of degradation of HMG-
CoA reductase, possibly by increasing the rate of transport of
the membrane-bound enzyme to the site of its degradation.

Lowering of serum cholesterol levels decreases the risk
of coronary heart disease. A number of drugs called statins
are potent competitive inhibitors of HMG-CoA reductase.
Statins are often used as part of the treatment of hypercho-
lesterolemia because they can effectively lower blood choles-
terol levels. Another useful approach is to bind bile salts in
the intestine to resin particles, to prevent their reabsorption.
More cholesterol must then be converted to bile salts. Inhibi-
tion of HMG-CoA reductase may not be the most desirable
method for controlling cholesterol levels because mevalonate
is needed for the synthesis of important molecules such as
ubiquinone.

► Structure of HMG-CoA
and two common statins.

3'-ADP

Figure 16.20 ►

Other products of isopentenyl diphosphate
and cholesterol metabolism.

Terpenes

(plant secondary <-
metabolites)
Gibberellins

Acetyl CoA

Isopentenyl

diphosphate

Quinones and phytol side
chain of chlorophyll

Vitamins A, E, K

Testosterone

/3-Estradiol

1,25-Dihydroxyvitamin D 3

16.6 Synthesis of Cholesterol

493

cholesterol. The two pathways for the biosynthesis of isopentyl diphosphate (Section
16.6A) are much more ancient than the more recent cholesterol biosynthesis pathway.

Cholesterol is the precursor of bile salts, which facilitate intestinal absorption of
lipids; vitamin D that stimulates Ca© uptake from the intestine; steroid hormones such
as testosterone and /3-estradiol that control sex characteristics; and steroids that control
salt balance. The principal product of steroid synthesis in mammals is cholesterol itself,
which modulates membrane fluidity and is an essential component of the plasma mem-
brane of animal cells.

16.7 Fatty Acid Oxidation

Fatty acids, released from triacylglycerols (Section 16.9), are oxidized by a pathway that
degrades them by removing two-carbon units at each step. The two-carbon fragments
are transferred to coenzyme A to form acetyl CoA, and the remainder of the fatty acid
re-enters the oxidative pathway. This degradative process is called the /3-oxidation path-
way because the /3-carbon atom (C-3) of the fatty acid is oxidized. Fatty acid oxidation
is divided into two stages: activation of fatty acids and degradation to two-carbon frag-
ments (as acetyl CoA). The NADH and ubiquinol (QH 2 ) produced by the oxidation of
fatty acids can be oxidized by the respiratory electron transport chain, and the acetyl
CoA can enter the citric acid cycle.

Acetyl CoA can be completely oxidized by the citric acid cycle to yield energy (in the
form of ATP) that can be used in other biochemical pathways. The carbon atoms from
fatty acids can also be used as substrates for amino acid synthesis since several of the in-
termediates in the citric acid cycle are diverted to amino acid biosynthesis pathways
(Section 13.6). In those organisms that possess a glyoxylate pathway (Section 13.7), acetyl
CoA from fatty acid oxidation can be used to synthesize glucose via the gluconeogenesis
pathway.

The oxidation of fatty acids occurs as part of the normal turnover of membrane
lipids. Thus, bacteria, protists, fungi, plants, and animals all have a /3-oxidation pathway.
In addition to its role in normal cellular metabolism, fatty acid oxidation is a major
component of fuel metabolism in animals. A significant percentage of dietary food con-
sists of membrane lipids and fat and this rich course of energy is exploited by oxidizing
fatty acids. In this section we describe the basic biochemical pathways of fatty acid oxi-
dation. In the following sections we will discuss the role of fatty acid oxidation in mam-
malian fuel metabolism.

A. Activation of Fatty Acids

The activation of fatty acids for oxidation is catalyzed by acyl-CoA synthetase (Figure 16.7).
This is the same activation step that is required for the synthesis of polyunsaturated fatty
acids and complex lipids.

B. The Reactions of /3-Oxidation

In eukaryotes, /3-oxidation takes place in mitochondria and in specialized organelles
called peroxisomes. In bacteria, the reactions take place in the cytosol. Four steps are re-
quired to produce acetyl CoA from fatty acyl CoA: oxidation, hydration, further oxida-
tion, and thiolysis (Figure 16.21). We focus first on the oxidation of a saturated fatty
acid with an even number of carbon atoms.

In the first oxidation step, acyl- CoA dehydrogenase catalyzes the formation of a
double bond between the C-2 and C-3 atoms of the acyl group forming trans 2-enoyl
CoA. There are several separate acyl-CoA dehydrogenase isozymes, each with a different
chain length preference: short, medium, long, or very long.

When the double bond is formed, electrons from fatty acyl CoA are transferred
to the FAD prosthetic group of acyl-CoA dehydrogenase and then to another FAD
prosthetic group bound to a mobile, water-soluble, protein coenzyme called electron

KEY CONCEPT

/^-Oxidation is an ancient and ubiquitous
pathway for degradation of fatty acids.

O O

ii ii

R — CH 2 — C— CH 2 — C— S-CoA

4 3 2 1

8 f3 a

3-ketoacyl CoA
(3-oxoacyl CoA)
(/3-ketoacyl CoA)

▲ 3-ketoacyl CoA, 3-oxoacyl CoA, /3-ketoacyl
CoA

▲ Bear bile. In Vietnam bears are kept in
captivity — often under deplorable conditions —
and bile is extracted from their stomachs on
a regular basis. Bear bile is thought to be an
effective remedy for fever and poor eyesight.

494 CHAPTER 16 Lipid Metabolism

-ch 2 — ch 2 — ch 2 -

3 2

S-CoA

R — CH 2 — C — CH 2 — C — S-CoA
3-Ketoacyl CoA

R — CH 2 — C = C — C — S-CoA
H

trans- A 2 -Enoyl CoA

H,0

I II

R — CH 2 — C — CH 2 — C — S-CoA
OH

L-3-Hydroxyacyl CoA

FADH

ETF

FAD

▲ Figure 16.21

/3-oxidation of saturated fatty acids. One round of /3-oxidation consists of four enzyme-catalyzed reac-
tions. Each round generates one molecule each of QH 2 , NADH, acetyl CoA, and a fatty acyl CoA
molecule two carbon atoms shorter than the molecule that entered the round. (ETF is the electron-
transferring flavoprotein, a water-soluble protein coenzyme.)

▲ Human medium chain acyl-CoA synthetase.

The products of the reaction, AMP and acyl
CoA, are bound in the active site. The en-
zyme is a dimer but only one subunit is
shown. [PDB 3EQ6]

transferring flavoprotein (ETF, Figure 16.22). (ETF also accepts electrons from several
other flavoproteins that are not involved in fatty acid metabolism.) Electrons are then
passed to Q in a reaction catalyzed by ETF: ubiquinone oxidoreductase. This enzyme is
embedded in the membrane and QH 2 from fatty acid oxidation enters the pool of QH 2
that can be oxidized by the membrane-associated electron transport system.

The second step is a hydration reaction. Water is added to the unsaturated trans 2-
enoyl CoA produced in the first step to form the L isomer of 3-hydroxyacyl CoA. The
enzyme is 2-enoyl-CoA hydratase.

The third step is a second oxidation catalyzed by L-3-hydroxyacyl-CoA dehydrogenase.
This production of 3-ketoacyl CoA from 3-hydroxyacyl CoA is an NAD® -dependent
reaction. The resulting reducing equivalents (NADH) can be used directly in biosynthe-
sis pathways or they can be oxidized by the membrane-associated electron transport
system.

Finally, in Step 4, the nucleophilic sulfhydryl group of HS-CoA attacks the carbonyl
carbon of 3-ketoacyl CoA in a reaction catalyzed by 3-ketoacyl- CoA thiolase. This enzyme,
also called thiolase II, is related to the acetoacyl-CoA thiolase (thiolase I) that we encountered

16.7 Fatty Acid Oxidation

495

▲ Figure 16.22

Model of the medium chain acyl-CoA dehydrogenase (MCAD) bound to ETF. The MCAD subunits are colored green and the ETF subunits are colored blue.
Bound FADs are represented as space-filling molecules (yellow). The model is based on the structure in PDB entry 2A1T containing a mutant protein
that blocks movement of the FAD domain of ETF. The left-hand side of the dimer shows the probable position of the FAD domain during transfer of
electrons from MCAD to ETF and the right-hand side shows the position of the FAD domain in free, unbound ETF. The flexibility of the FAD domain
as it shifts from one position to another is responsible for its lack of resolution in the wild-type ETF: MCAD crystal structure. (Toogood et al., 2004;
Toogood et al., 2005)

in the isopentenyl diphosphate pathway (Section 16.6A). Acetoacyl-CoA thiolase is specific
for acetoacetyl Co A, while 3-ketoacyl-CoA thiolase acts on long chain fatty acid derivatives.
The release of acetyl CoA leaves a fatty acyl CoA molecule shortened by two carbons. This
acyl CoA molecule is a substrate for another round of the four reactions and the metabolic
spiral continues until the entire molecule has been converted to acetyl CoA.

As the fatty acyl chain becomes shorter, the first step is catalyzed by acyl- CoA dehy-
drogenase isozymes with preferences for shorter chains. Interestingly, the first three reac-
tions of fatty acid oxidation are chemically parallel to three steps of the citric acid cycle.
In these reactions, an ethylene group ( — CH 2 CH 2 — , as in succinate) is oxidized to a
two-carbon unit containing a carbonyl group ( — COCH 2 — , as in oxalo acetate). The
steps are the reverse of the reactions in the fatty acid synthesis pathway (Section 16. 1C).

In eukaryotes, fatty acid oxidation also occurs in peroxisomes. In fact, peroxisomes are
the only site of fatty acid /3-oxidation in most eukaryotes (but not mammals). In peroxi-
somes, the initial oxidation step is catalyzed by acyl- CoA oxidase — an enzyme that is
homologous to the acyl-CoA dehydrogenease that catalyzes the first oxidation in mitochon-
dria. The peroxisomal enzyme transfers electrons to 0 2 to form hydrogen peroxide (H 2 0 2 ).

Fatty acyl CoA + 0 2 > trans- A 2 -Enoyl CoA + H 2 0 2 (16.7)

In bacterial and mitochondrial /3- oxidation the product of the first oxidation step is
QH 2 that can be used in the respiratory electron transport chain. This results in synthesis of
ATP — each QH 2 molecule is equivalent to 1.5 molecules of ATP (Section 14.11). There is no
membrane-associated electron transport system in peroxisomes and this is why a different
type of oxidation-reduction takes place in peroxisomes. It also means that fewer ATP equiv-
alents are produced during peroxisomal /3-oxidation. In mammals, where both mitochon-
drial and peroxisomal pathways exist, the peroxisomal /3- oxidation pathway degrades very
long chain fatty acids, branched fatty acids, long chain dicarboxylic acids, and possibly

▲ Peroxisomes. Indian Muntjac ( Muntiacus
muntjak) fibroblast cells were stained with
green reagent to show peroxisomes. Actin
fibers are stained red and nuclear DNA is
purple. The small peroxisomes are scattered
throughout the cytoplasm. [http://www.
m i c rosco py u . co m/st at i cga llery/
fluorescence/muntjac.html]

496 CHAPTER 16 Lipid Metabolism

Fatty acid synthesis

Acyl ACP (C n + 2 )

NADP©

Reduction

NADPH + H©-

trans-A 2 -Enoyl ACP (C n + 2 )

Dehydration

D-3-Hydroxyacyl ACP (C n + 2 )

NADP©-

Reduction

NADPH + H©-

3-Ketoacyl ACP (C n + 2 )
HS-ACP + C0 2 «^

Malonyl ACP •

Acyl ACP (C n )

Condensation

/3-oxidation
Acyl CoA (C n + 2 )

trans unsaturated fatty acids producing smaller, more polar compounds that can be ex-
creted. Most of the common fatty acids are degraded in mitochondria.

C. Fatty Acid Synthesis and /3-Oxidation

Fatty acid synthesis involves carbon-carbon bond formation (condensation) followed
by reduction, dehydration, and reduction steps in preparation for the next condensa-
tion reaction. The reverse reactions — oxidation, hydration, oxidation, and carbon-carbon
bond cleavage — are part of the degradation pathway of /3- oxidation. We compare the
two pathways in Figure 16.23.

The active thioesters in fatty acid oxidation are CoA derivatives whereas the inter-
mediates in fatty acid synthesis are bound as thioesters to acyl carrier protein (ACP). In
both cases, the acyl groups are attached to phosphopantetheine. Synthesis and degrada-
tion both proceed in two-carbon steps. However, oxidation results in a two-carbon
product, acetyl CoA, whereas synthesis requires a three- carbon substrate, malonyl ACP
that transfers a two-carbon unit to the growing chain releasing C0 2 . Reducing power for
synthesis is supplied by NADPH, whereas oxidation depends on NAD® and ubiquinone
(via the electron-transferring flavoprotein). Finally, the intermediate in fatty acid syn-
thesis is D-3-hydroxyacyl-ACP whereas the L isomer (L-3-hydroxyacyl-CoA) is pro-
duced during /3-oxidation.

The biosynthesis and catabolic pathways are catalyzed by a completely different set
of enzymes and the intermediates form separate pools due to the fact that they are
bound to different cofactors (CoA and ACP). In eukaryotic cells the two opposing path-
ways are physically separated. The biosynthesis enzymes are found in the cytosol and
the 13 - oxidation enzymes are confined to mitochondria and peroxisomes.

Oxidation

>QH 2

frans-A 2 -Enoyl CoA (C n + 2 )

Hydration

L-3-Hydroxyacyl CoA (C n + 2 )
f NAD©

Oxidation

^NADH + H 0
3-Ketoacyl CoA (C n + 2 )
-HS-CoA

Acetyl CoA
Acyl CoA (C n )

Thiolysis

▲ Figure 16.23

Fatty acid synthesis and /3-oxidation.

Figure 16.24 ►

Carnitine shuttle system for transporting fatty
acyl CoA into the mitochondrial matrix. The

path of the acyl group is traced in red.

D. Transport of Fatty Acyl CoA into Mitochondria

Long-chain fatty acyl CoA formed in the cytosol cannot diffuse across the inner mito-
chondrial membrane into the mitochondrial matrix where the reactions of /3- oxidation
occur in mammals. A transport system, called the carnitine shuttle system, actively
transports fatty acids into mitochondria (Figure 16.24). In the cytosol, the acyl group of

0N(CH 3 ) 3

L-Carnitine

©N(CH 3 ) 3

Acylcarnitine

Fatty acyl CoA

16.7 Fatty Acid Oxidation 497

BOX 16.4 A TRIFUNCTIONAL ENZYME FOR /3-OXIDATION

NAD

Many species contain a trifunctional en-
zyme for /3-oxidation. The 2-enoyl-CoA
hydratase (ECH) and L-3-hydroxyacyl-
CoA dehydrogenase (HACD) activities
are located on a single polypeptide chain
(a subunit). The 3-ketoacyl-CoA thiolase
(KACT) activity is localized to the /3 sub-
unit and the two subunits combine to
form a protein with a 2 & 2 quaternary
structure.

The structure of a bacterial enzyme is
shown in the figure. During /3-oxidation
the product of the first reaction, trans-2-
enoyl CoA, binds to the ECH site of the
trifunctional enzyme. The substrate then
undergoes the next three reactions within
the cavity formed by the ECH, HACD, and
KACT active sites in each half of the
dimer. The two intermediates in the path-
way are not released during these reac-
tions because they are bound by their CoA
termini. This is an example of metabolic
channeling by a multienzyme complex.

▲ Structure of the fatty acid /3-oxidation multienzyme complex from the bacterium Pseudomonas

fragi. In this structure a molecule of acyl CoA is bound at each of the KACT sites. [PDB 1WDK]

fatty acyl CoA is transferred to the hydroxyl group of carnitine to form acylcarnitine in
a reaction catalyzed by carnitine acyltransferase I, also called carnitine palmitoyltrans-
ferase I (CPTI). The enzyme is associated with the outer membrane of the mitochondria.

This reaction is a key site for regulation of the oxidation of intracellular fatty acids. The
acyl ester acylcarnitine is a “high energy” molecule with a free energy of hydrolysis similar
to that of a thiol ester. Acylcarnitine then enters the mitochondrial matrix in exchange for
free carnitine via the carnitine: acylcarnitine translocase. In the mitochondrial matrix, the
isozyme carnitine acyltransferase II catalyzes the reverse of the reaction catalyzed by carni-
tine acyltransferase I. The effect of the carnitine shuttle system is to remove fatty acyl CoA
from the cytosol and regenerate fatty acyl CoA in the mitochondrial matrix.

The carnitine shuttle system is not used in most eukaryotes since fatty acid oxida-
tion takes place in the peroxisomes. Fatty acids are transported into peroxisomes by a
different mechanisms of course, no transport mechanism is required in prokaryotes
since all these reactions take place in the cytoplasm.

KEY CONCEPT

Unlike the pathways for gluconeogenesis
and glycolysis, the pathways for the
synthesis and degradation of fatty acids
are completely different.

In Section 16.7D we compare the cost
of fatty acid synthesis to the energy
recovered in /?-oxidation.

E. ATP Generation from Fatty Acid Oxidation

The complete oxidation of fatty acids supplies more energy than the oxidation of an
equivalent amount of glucose. As is the case in glycolysis, the energy yield of fatty acid
oxidation can be estimated from the total theoretical yield of ATP (Section 13.5). As an
example, let’s consider the balanced equation for the complete oxidation of one mole-
cule of stearate (C 18 ) by eight cycles of /3-oxidation. Stearate is converted to stearoyl
CoA at a cost of two ATP equivalents and the oxidation of steroyl CoA yields acetyl CoA
and the reduced coenzymes QH 2 and NADH.

Stearoyl CoA + 8 HS-CoA + 8 Q + 8 NAD® >

9 Acetyl CoA + 8 QH 2 + 8 NADH + 8 H®

(16.8)

498 CHAPTER 16 Lipid Metabolism

We can calculate the theoretical yield of 9 molecules of acetyl CoA by assuming that they
enter the citric acid cycle where they are completely oxidized to C0 2 . These reactions
produce 10 ATP equivalents for each molecule of acetyl CoA. The net yield from oxida-
tion of stearate is 120 ATP equivalents.

Eight cycles of / 3 -oxidation yield

8 QH 2 « 12 ATP

8NADH « 20 ATP

9 molecules of acetyl CoA ~ 90 ATP

activation of stearate ~ —2 ATP

Total = 120 ATP

HCO 3 0

Bicarbonate

H O

I II

H — C — CH 2 — C — S-CoA
H

Propionyl CoA

ATP — ^

(Biotin)
Pi + ADP^

Propionyl-CoA

carboxylase

H — C— H

O 1

e OOC— c — c — S-CoA

I II

H O

D-Methylmalonyl CoA

Methylmalonyl-CoA

racemase

By comparison, the oxidation of glucose to C0 2 and water yields approximately 32
ATP molecules. Since stearate has 18 carbons and glucose has only six carbons, we nor-
malize the yield of ATP from glucose by comparing the oxidation of three molecules of
glucose: 3 X 32 = 96 ATP. This theoretical ATP yield is only 80% of the value for
stearate. Fatty acids provide more energy per carbon atom than carbohydrates because
carbohydrates are already partially oxidized. Furthermore, because fatty acid moieties
are hydrophobic, they can be stored in large quantities as triacylglycerols without large
amounts of bound water, as are found with carbohydrates. Anhydrous storage allows far
more energy to be stored per gram.

We can also calculate the cost of synthesizing stearate in order to compare it to the
energy recovered during /3-oxidation. For this calculation we need to know the cost of
synthesizing acetyl CoA from C0 2 . This value (17 ATP equivalents) is obtained from
the reactions of C0 2 fixation in plants (Section 15.4C).

8 acetyl CoA —> 8 malonyl ACP

8 synthesis steps 16 NADPH

9 acetyl CoA 9 X 17

Total

8 ATP
40 ATP
153 ATP
201 ATP

The energy recovered in the degradation of stearate is about 60% (120/201) of the total
theoretical energy required for its synthesis. This is a typical example of biochemical
efficiency.

H O

O 1 11

e OOC — C — C — S-CoA

H — C — H

L-Methylmalonyl CoA

(Adenosyl- Methylmalonyl-CoA
cobalamin) mutase

O 1

^OOC — C — H

H — c— : — S-CoA

I II

H O

Succinyl CoA
▲ Figure 16.25

Conversion of propionyl CoA to succinyl CoA.

F. /3-Oxidation of Odd-Chain and Unsaturated Fatty Acids

Most fatty acids have an even number of carbon atoms. Odd-chain fatty acids are syn-
thesized by bacteria and by some other organisms. Odd-chain fatty acids are oxidized by
the same sequence of reactions as even- chain fatty acids except that the product of the
final thiolytic cleavage is propionyl CoA (CoA with a C 3 acyl group) rather than acetyl
CoA (CoA with a C 2 acyl group). In mammals, propionyl CoA can be converted to suc-
cinyl CoA in a three step pathway (Figure 16.25).

The first reaction is catalyzed by propionyl- CoA carboxylase, a biotin-dependent en-
zyme that incorporates bicarbonate into propionyl CoA to produce D-methylmalonyl
CoA. Methylmalonyl-CoA racemase catalyzes the conversion of D-methylmalonyl CoA to
its L isomer. Finally, methylmalonyl-CoA mutase catalyzes the formation of succinyl CoA.

Methylmalonyl-CoA mutase is one of the few enzymes that require adenosylcobal-
amin as a cofactor. We learned in Section 7.12 that adenosylcobalamin-dependent
enzymes catalyze intramolecular rearrangements in which a hydrogen atom and a sub-
stituent on an adjacent carbon atom exchange places. In the reaction catalyzed by
methylmalonyl-CoA mutase, the — C(O) — S-CoA group exchanges with a hydrogen
atom of a methyl group (Figure 7.28).

The succinyl CoA molecule formed by the action of methylmalonyl-CoA mutase is
metabolized to oxaloacetate. Since oxaloacetate is a substrate for gluconeogenesis, the

16.7 Fatty Acid Oxidation 499

Linoleoyl CoA

(18:2, c/'s, c/s- A 9,12 )

(12:2, c/'s, c/s- A 3 6 )

H H H H

-C = C— CH 2 — C = C — ch 2

CD cz>^ Three rounds of /3-oxidation

c=x>

I o

H H
C = C

4 3 2

H H

-c=c— ch 2 — c=c— ch 2 — c— S-CoA

A 3 , A-Enoyl-CoA isomerase

(12:2, tans.cis- A 26 )

H H H

-C = C— CH 2 — CH 2 — C = C — c— S-CoA

(D ciD One round of /3-oxidation

(10:1, c/s- A 4 )

(10:2, tans.cis- A 24 )

(10:1, tans- A 3 )

(10:1, fans-A 2 )

H H II

-C=C—CH 2 — CH 2 —C— S-CoA

Acyl-CoA dehydrogenase
(first reaction of /3-oxidation)

H H H II

-C = C — C = C— C— S-CoA

, — NADPH,H®

( 5 ) 2,4-Dienoyl-CoA reductase

©>NADP®

H II

-CH 2 — C = C — CH 2 — c— S-CoA

A 3 , A 2 -Enoyl-CoA isomerase
(same enzyme as Step 2)

H II

-CH,— CH,— C = C — C— S-CoA

One round of /3-oxidation

-CH 2 —CH 2 — C — S-CoA

Figure 16.26 ►

Oxidation of linoleoyl CoA. Oxidation requires
two enzymes: enoyl-CoA isomerase and
2,4-dienoyl-CoA reductase — in addition to
the enzymes of the /3-oxidation pathway.

propionyl group derived from the (5 - oxidation of an odd- chain fatty acid can be con-
verted to glucose.

The oxidation of unsaturated fatty acids requires two enzymes in addition to those
usually needed for the oxidation of saturated fatty acids. The oxidation of the Coenzyme A
derivative of linoleate (18:2 cis,cis A 9,12 -octadecadienoate) illustrates the modified path-
way (Figure 16.26).

500 CHAPTER 16 Lipid Metabolism

▲ The camel’s hump stores fat for energy pro-
duction when food is scarce. The hump of the
camel contains fat that is used to supply en-
ergy. It does not store water. The ability of
camels to go for long periods of time without
water is due to completely different adapta-
tions having nothing to do with fat metabo-
lism. The camel shown here is the Arabian
camel or dromedary, Camelus dromedarius.

Like all polyunsaturated fatty acids linoleoyl CoA has both odd-numbered and
even-numbered double bonds (its double bonds are separated by a methylene group).
Unsaturated fatty acids are normal substrates for the enzymes of the /3-oxidation path-
way until an odd-numbered double bond of the shortened fatty acid chain interferes
with catalysis. In this example, three rounds of /3- oxidation convert linoleoyl CoA to the
C 12 molecule 12:2 ds,ds- A 3,6 -dienoyl CoA (step 1). This molecule has a ds- 3,4 double
bond rather than the usual trans- 2,3 double bond that would be produced during 13-
oxidation of saturated fatty acids. The cis- 3,4 intermediate is not a substrate for
2-enoyl-CoA hydratase since the normal /3-oxidation enzyme is specific for trans acyl
CoAs and, in addition, the double bond is in the wrong position for hydration.

The inappropriate double bond is rearranged from A 3 to A 2 to produce the C 12
molecule 12:2 trans, cis- A 2,6 - dienoyl CoA in a reaction catalyzed by A 3 ,A 2 -enoyl-CoA
isomerase (step 2). This product can re-enter the /3-oxidation pathway and another
round of /3-oxidation can be completed resulting in the C 10 molecule 10:1 cis- A 4 -enoyl
CoA (step 3). The first enzyme of the /3- oxidation pathway, acyl- CoA dehydrogenase,
acts on this compound, producing the C 10 molecule 10:2 trans, cis- A 2,4 -dienoyl CoA.
This resonance-stabilized diene resists hydration. 2,4-Dienoyl-CoA reductase catalyzes
the NADPH-dependent reduction of the diene (step 5) to produce a C 10 molecule with
a single double bond (10:1 trans- A 3 -enoyl CoA). This product (like the substrate in
step 2) is acted on by A 3 , A 2 -enoyl-CoA isomerase to produce a compound that contin-
ues through the /3-oxidation pathway. Note that the isomerase can convert both cis- A 3
and trans- A 3 double bonds to the trans- A 2 intermediate.

The oxidation of a monounsaturated fatty acid with a cis double bond at an odd-
numbered carbon (e.g., oleate) requires the activity of the isomerase but not the re-
ductase, in addition to the enzymes of /3-oxidation. Oleoyl (18:1 cis- A 3 ) CoA undergoes
three cycles of /3-oxidation, forming three molecules of acetyl CoA and the CoA ester
of the (12:1 ds-A 3 ) acid. A 3 , A 2 -Enoyl-CoA isomerase then catalyzes conversion
of the 12-carbon enoyl CoA to a 12-carbon trans- A 2 enoyl CoA, which can undergo
/3-oxidation.

▲ Myelin sheath. These nerve fibers are
coated with several layers of myelin mem-
branes (colored purple) forming a protective
sheath around the axons. Plasmalogens are
important components of myelin mem-
branes. The symptoms of multiple sclerosis
(MS) are caused by degradation of myelin in
the brain and spinal cord leading to loss of
motor control.

16.8 Eukaryotic Lipids Are Made at
a Variety of Sites

Eukaryotic cells are highly compartmentalized. The compartments can have quite dif-
ferent functions, and their surrounding membranes can have quite distinct phospho-
lipid and fatty acyl constituents. Most lipid biosynthesis in eukaryotic cells occurs in the
endoplasmic reticulum. Phosphatidylcholine, phosphatidylethanolamine, phos-
phatidylinositol, and phosphatidylserine, for example, are all synthesized in the ER. The
biosynthesis enzymes are membrane bound with their active sites oriented toward the
cytosol so that they have access to the water-soluble cytosolic compounds. The major
phospholipids are incorporated into the ER membrane. From there they are trans-
ported to other membranes in the cell in vesicles that travel between the endoplasmic
reticulum and Golgi apparatus and between the Golgi apparatus and various mem-
brane target sites. Soluble transport proteins also participate in carrying phospholipids
and cholesterol to other membranes.

Although the endoplasmic reticulum is the principal site of lipid metabolism in the
cell, there are also lipid-metabolizing enzymes at other locations. For instance, mem-
brane lipids can be tailored to give the lipid profile characteristic of individual cellular
organelles. In the plasma membrane, acyltransferase activities catalyze the acylation of
lysophospholipids. Mitochondria have the enzyme phosphatidylserine decarboxylase
that catalyzes the conversion of phosphatidylserine to phosphatidylethanolamine.
Mitochondria also contain the enzymes responsible for the synthesis of diphosphatidyl-
glycerol (cardiolipin, Table 9.2), a molecule found uniquely in the inner membrane of
the mitochondrion. Lysosomes possess various hydrolases that degrade phospholipids
and sphingolipids. Peroxisomes possess enzymes involved in the early stages of ether

16.8 Eukaryotic Lipids Are Made at a Variety of Sites 501

lipid synthesis. Defects in peroxisomal formation can lead to poor plasmalogen synthe-
sis, with potentially fatal consequences.

The tissues of the central nervous system are especially prone to damage. In those
tissues plasmalogens constitute a substantial portion of the lipids of the myelin sheath.
Often, different subcellular locations have a different set of enzymes (isozymes) respon-
sible for the biosynthesis of different, segregated pools of lipids, with each pool having
its own biological function.

16.9 Lipid Metabolism is Regulated by
Hormones in Mammals

Fatty acids are no longer oxidized in mitochondria when the energy supply is sufficient
to meet the immediate needs of an organism. Instead, they are transported to adipose
tissue where they are stored for future use when energy is needed (e.g., lack of food).
This aspect of lipid metabolism is similar to the strategy in carbohydrate metabolism
where excess glucose is stored in specialized cells as glycogen (animals) or starch
(plants).

The mobilization and storage of lipids requires communication between different
tissues. Hormones that circulate in the blood are ideally suited to act as signals between
cells. Lipid metabolism must be coordinated with carbohydrate metabolism, so it’s not
surprising that the same hormones also affect the synthesis, degradation, and storage of
carbohydrates.

Glucagon, epinephrine, and insulin are the principal hormonal regulators of fatty
acid metabolism. Glucagon and epinephrine are present in high concentrations in the
fasted state and insulin is present in high concentrations in the fed state. The concentra-
tion of circulating glucose must be maintained within fairly narrow limits at all times.
In the fasted state, carbohydrate stores become depleted and synthesis of carbohydrates
must occur to maintain the level of glucose in the blood. To further relieve pressure on
the limited supply of glucose, fatty acids are mobilized to serve as fuel, and many tissues
undergo regulatory transitions that decrease their use of carbohydrates and increase
their use of fatty acids. The opposite occurs in the fed state when carbohydrates are used
as fuel and precursors for fatty acid synthesis.

The key regulatory enzyme for fatty acid synthesis is acetyl- CoA carboxylase. High
insulin levels after a meal inhibit the hydrolysis of stored triacylglycerols and stimulate
the formation of malonyl CoA by acetyl-CoA carboxylase. Malonyl CoA allosterically
inhibits carnitine acyltransferase I. As a result, fatty acids remain in the cytosol rather
than being transported into mitochondria for oxidation. Regulation of fatty acid syn-
thesis and degradation is reciprocal, with increased metabolism by one pathway bal-
anced by decreased activity in the opposing pathway. In animals this regulation is
achieved by hormones that indirectly affect the activities of the enzymes.

Triacylglycerols are delivered to adipose tissue in the form of lipoproteins that cir-
culate in blood plasma (Section 16.10B). When they arrive at adipose tissue the triglyc-
erols are hydrolyzed to release fatty acids and glycerol that are then taken up by
adipocytes. Hydrolysis is catalyzed by lipoprotein lipase (LPL), an extracellular enzyme
bound to endothelial cells of the capillaries of adipose tissue. Following entry into
adipocytes, the fatty acids are re-esterified for storage as triacylglycerols.

Subsequent mobilization, or release, of fatty acids from adipocytes depends on
metabolic needs. A hormone-sensitive lipase in adipocytes catalyzes the hydrolysis of
triacylglycerols to free fatty acids and monoacylglycerols. Although hormone-sensitive
lipase can also catalyze the conversion of monoacylglycerols to glycerol and free fatty
acids, a more specific and more active monoacylglycerol lipase probably accounts for
most of this catalytic activity.

The hydrolysis of triacylglycerols is inhibited in the fed state by high concentrations
of insulin. When carbohydrate stores are depleted and insulin concentrations are low,
an increased concentration of epinephrine stimulates triacylglycerol hydrolysis. Epi-
nephrine binds to the /3-adrenergic receptors of adipocytes leading to activation of the

Hormone signaling pathways are
described in Section 9.12.

502 CHAPTER 16 Lipid Metabolism

BOX 16.5 LYSOSOMAL STORAGE DISEASES

There are no metabolic diseases associated with defects in the
sphingolipid biosynthesis pathways. It is likely that mutations
in the genes for biosynthesis enzymes are lethal since sphin-
golipids are essential membrane components. In contrast,
defects in the sphingolipid degradation pathway can have se-
rious clinical consequences. Sphingolipid catabolism is largely
carried out in the lysosomes of cells. Lysosomes contain a va-
riety of glycosidases that catalyze the stepwise hydrolytic
removal of sugars from the oligosaccharide chains of sphin-
golipids. There are certain inborn errors of metabolism in
which a genetic defect leads to a deficiency in a particular
degradative lysosomal enzyme resulting in lysosomal storage
diseases. The accumulation of nondegradable lipid by-products
can cause lysosomes to swell leading to cellular and ultimately
tissue enlargement. This is particularly deleterious in central
nervous tissue that has little room for expansion. Swollen

lysosomes accumulate in the cell bodies of nerve cells and
lead to neuronal death, possibly by leakage of lysosomal en-
zymes into the cell. As a result, blindness, mental retardation,
and death can occur. In Tay-Sachs disease, for instance, there
is a deficiency in hexosaminidase A, which catalyzes the re-
moval of N - acetylgalactosamine from the oligosaccharide
chain of gangliosides. If removal of this sugar does not occur,
the disassembly of gangliosides is blocked, leading to a
buildup of the nondegradable by-product, ganglioside G M2 .
(The complete structure of ganglioside G M2 is shown in
Figure 9.12.)

Schematic pathways for the formation and degradation
of a variety of sphingolipids are shown in the accompanying
figure. A number of defects in sphingolipid metabolism,
whose clinical manifestations are termed sphingolipidoses , are
identified there.

Sphingosine

Glucocerebroside

Trihexosylceramide

Globoside

(Galy— [ Cer
0 °3 S Sulfatide

Disease

Mental

retardation

Liver

damage

Myelin

defects

Specialized symptoms

Fatal

Farber's

Damage to joints, granulomas

Niemann-Pick

Gaucher's

Bone damage

Frequently

Krabbe's

Globoid bodies in brain

Fabry's

Rash, kidney failure

Metachromatic

leukodystrophy

Paralysis, dementia

Tay-Sachs

Blindness, seizures

Sandhoff's

Same as Tay-Sachs;
progresses more rapidly

Generalized

gangliosidosis

Bone damage

16.9 Lipid Metabolism is Regulated by Hormones in Mammals 503

Epinephrine

▲ Figure 16.27

Triacylglycerol degradation in adipocytes. Epinephrine initiates the activation of protein kinase A, which catalyzes the phosphorylation and activation
of hormone-sensitive lipase. The lipase catalyzes the hydrolysis of triacylglycerols to monoacylglycerols and free fatty acids. The hydrolysis of monoa-
cylglycerols is catalyzed by monoacylglycerol lipase.

cAMP- dependent protein kinase A. Protein kinase A catalyzes the phosphorylation and
activation of hormone-sensitive lipase (Figure 16.27).

Glycerol and free fatty acids diffuse through the adipocyte plasma membrane and enter
the bloodstream. Glycerol is metabolized by the liver, where most of it is converted to glu-
cose via gluconeogenesis. Free fatty acids are poorly soluble in aqueous solution and travel
through blood bound to serum albumin (Section 16.9C). Fatty acids are carried to tissues
such as heart, skeletal muscle, and liver, where they are oxidized in mitochondria to release
energy. Fatty acids are a major source of energy during the fasting state (e.g., while we sleep).

At the same time, an increase in glucagon levels inactivates acetyl-CoA carboxylase,
the enzyme that catalyzes the synthesis of malonyl CoA in the liver. The result is in-
creased transport of fatty acids into mitochondria and greater flux through the
/3-oxidation pathway. The high concentrations of acetyl CoA and NADH that are pro-
duced by fatty acid oxidation decrease glucose and pyruvate oxidation by inhibiting the
pyruvate dehydrogenase complex. Thus, not only are fatty acid oxidation and storage
reciprocally regulated but fatty acid metabolism is also regulated so that storage is fa-
vored in times of plenty (such as immediately after feeding) and fatty acid oxidation
proceeds when glucose must be spared.

Citrate — a precursor of cytosolic acetyl CoA — activates acetyl- CoA carboxylase in
vitro , but the physiological relevance of this activation has not been fully established.
Acetyl- CoA carboxylase is inhibited by fatty acyl CoA. The ability of fatty acid deriva-
tives to regulate acetyl- CoA carboxylase is physiologically appropriate; an increased
concentration of fatty acids causes a decrease in the rate of the first committed step of
fatty acid synthesis. Acetyl CoA- carboxylase activity is also under hormonal control.
Glucagon stimulates phosphorylation and concomitant inactivation of the enzyme in
the liver, and epinephrine stimulates its inactivation by phosphorylation in adipocytes.
Several protein kinases can catalyze phosphorylation and thus inhibition of acetyl- CoA
carboxylase. The action of AMP activated protein kinase inactivates both fatty acid syn-
thesis (by inhibiting the acetyl- CoA carboxylase step) and steroid synthesis in the pres-
ence of a high AMP/ATP ratio.

LIPID METABOLISM

GUT

BIOSYNTHESIS

I LIVER) Celt Membrane
Cytosol

FATTY ACID SYNTHASE

{ 2 . 3 , 1 .86)

Muftr-enzyme com plex includes all the enzymes of
Fatly Acid Synthesis except Acetyl-CoA carboxylase

CHjOOO-

CHjCtOHlCHjCOSGaA'

3-0 H -3- methytgl y tary 1-CoA { H MG )

Committed
to Cholesterol
Biosynthesis

CHjGOCHa cH^oCHa&ocr ch3C«ioh>ch 2 oqo

Acetone-^-^Acetoacetate ^ 3-OH - Buty rate

Mitochondrial inner memtuarte
! KETONE BODIES \

Brain

Skeletal & Heart Muscle

Renal Cortex

(In starvation)^

ANABOLISM

CATABOLISM

8CH 3 CO.SCoA+ 14 NADPH + 14 H++ 7 ATP
Acetyl-CoA

GH 3 [CH 2 CH 2 ] 7 CO,SCoA
Patmitoyl-CoA

^ CH 3 [CH 2 CH 2 ] 7 C 0 SCqA+ 1 4 NAD P * + 7 BSC oA + 7 ADP+ 7 Pj

Palmitoyl-CoA

/Ubiquinone + 7 H.O + 7NAD*'+ 7 HSCoA — 8CH3CO .SCoA +7 Ubiquinol + 7 NADH + 7 H +

Acetyl-CoA

COMPLETE (AEROBIC) OXIDATION OF PALMITOYL CoA

CH 3 [CH 2 CH 2 ] 7 CO.SCoA + 2302 + - (106 AD P + 106Pf)

. I 6 CO 2 +119 H 2 D + HSCoA + -106 ATP

This is a fascinating equation which explains how some animals, such as camels and polar bears can survive in the most adverse environments
They can use fat ,not only as the sole source of energy, but also of water The killer whale cannot utilise sea-water but creates its own from fat

1.1.1

1 . 1/1

1.1.1

1.1.1.

1 . 1 . 1 .

1.1.1

12.4.

Glycerol-3- P-dehydrogenase J
HMG-CoA reductase
3-OH-acy 1-CoA dehydrogenase '■
Malatedehydrogenase «

M a late dehydrogenase 2

(oxaloacetate z

3-Oxoacy1-[ACP] f

3-OH-bulyryl-CoA |

Long-chain 3-OH-acyl-CcA £
Pyruvafedehydrogenase 2

3.1.10
.3.99.2
3.99 3
:.3.17
.3 19
',3.1.15
:.3.1.16
.3.120
.3.1.38
.3.1 39

ENZYMES

Enoyl-f AC P]-reductase
Butyryl-CoA dehydrogenase
Acyl-CoA dehydrogenase
Ca rn Itl ne-O-acy lira nsferase
Acelyl-CoA-C-acetyl transferase
Glycerol-3-P O-acyl transferase
Acetyl-CoA C-acy lira nsferase
Diacylglyee rolO-acyl iransferase
[ACPlS-acyl transferase
[AGP]S-malonyl transferase

2.3.1.51 l-Acylglycerol-3-P O-acyl
transferase

3-113 Triacylglycerol lipase
3.1.123 Acylglycerol lipase
3.1.123 Acylca mi tin e hydrolase
3. 1 1 34 Lipoprotein lipase
31,3.4 Phosphatidate phosphatase
4 1.1.4 Acetoaceiate decarboxylase
4.11.9 Malonyl-CoAdecarboxylase
4.1 34 OH-Methylglutaryl-CoA lyase

4 1.3,5 OH-Methylglutaryl-CoA synthase
4.13.7 Citrate sy nma se
4.13.3 ATPCitrafe lyase
4.2.1,17 Enoyl-CoA hydratase
4.2.1 .55 3- OH-ButyryLCq A dehydratase
4 2 1.58 C rotonyl-FAC P] hydratase
4,2.159 3-OH-octa noy I- 7

5 ] hydratase
l-[ACP] dehydratase
4-[ ACPI dehydratase

4.2. 1 ,61 3-OH-Palmitoyl-[ACPl dehyc
6.21.3 Long-chain-fatty -acid-Co A li
$4.11 Pyruvate carboxylase
6.4.12 Acetyl-CoA ca rPoxyla se

504

16.10 Absorption and Mobilization of Fuel Lipids 505

16.10 Absorption and Mobilization of Fuel Lipids

The fatty acids and glycerol that mammals use as metabolic fuels are obtained from tri-
acylglycerols in the diet and from adipocytes. The fats stored in adipocytes include fats
synthesized from the catabolism of carbohydrates and amino acids. Free fatty acids
occur only in trace amounts in cells — this is fortunate because, as anions, they are de-
tergents and at high concentrations could disrupt cell membranes. We begin our study
of lipid metabolism by examining the dietary uptake, transport, and mobilization of
fatty acids in mammals.

A. Absorption of Dietary Lipids

Most lipids in the diets of mammals are triacylglycerols with smaller amounts of phos-
pholipids and cholesterol. The digestion of dietary lipids occurs mainly in the small in-
testine, where suspended fat particles are coated with bile salts (Figure 16.28). Bile salts
are amphipathic cholesterol derivatives synthesized in the liver, collected in the gallblad-
der, and secreted into the lumen of the intestine. Micelles of bile salts solubilize fatty
acids and monoacylglycerols so that they can diffuse to and be absorbed by the cells of
the intestinal wall. Lipids are transported through the body as complexes of lipid and
protein known as lipoproteins.

Triacylglycerols are broken down in the small intestine by the action of lipases.
These enzymes are synthesized as zymogens in the pancreas and secreted into the small
intestine where they are activated. Pancreatic lipase catalyzes hydrolysis of the primary
esters (at C-l and C-3) of triacylglycerols releasing fatty acids and generating monoa-
cylglycerols (Figure 16.29). A small protein called colipase helps bind the water-soluble
lipase to the lipid substrates. Colipase also activates lipase by holding it in a conforma-
tion with an open active site. The fatty acids derived from dietary triacylglycerols are
primarily long chain molecules.

Most of these bile salts recirculate through the lower parts of the small intestine,
the hepatic portal blood, and then the liver. Bile salts circulate through the liver and in-
testine several times during the digestion of a single meal. Fatty acids are converted to
fatty acyl CoA molecules within the intestinal cells. Three of these molecules can com-
bine with glycerol, or two with a monoacylglycerol, to form a triacylglycerol. As described
below, these water- insoluble triacylglycerols combine with cholesterol and specific pro-
teins to form chylomicrons for transport to other tissues.

The fate of dietary phospholipids is similar to that of triacylglycerols. Pancreatic
phospholipases secreted into the intestine catalyze the hydrolysis of phospholipids
(Figure 9.8), which aggregate in micelles. The major phospholipase in the pancreatic secre-
tion is phospholipase A 2 , which catalyzes hydrolysis of the ester bond at C-2 of a glyc-
erophospholipid to form a lysophosphoglyceride and a fatty acid (Figure 16.30). A model
of phospholipase A 2 with a lipid substrate is shown in Figure 16.31. Lysophosphoglycerides
are absorbed by the intestine and re-esterified to glycerophospholipids in intestinal cells.

Lysophosphoglycerides are normally present in cells only at low concentrations. High
concentrations can disrupt cellular membranes by acting as detergents. This occurs, for
example, when snake venom phospholipase A 2 acts on phospholipids in red blood cells,
causing lysis of erythrocyte membranes. This is probably what killed Cleopatra.

Unlike other types of dietary lipids, most dietary cholesterol is unesterified. Dietary
cholesteryl esters are hydrolyzed in the lumen of the intestine by the action of an esterase.
Free cholesterol, which is insoluble in water, is solubilized by bile-salt micelles for ab-
sorption. Most cholesterol reacts with acyl CoA to form cholesteryl esters (Figure 9.16)
in the intestinal cells.

.SO*

coo 0

▲ Figure 16.28

Bile salts. The cholesterol derivatives tauro-
cholate and glycocholate are the most abun-
dant bile salts in humans. Bile salts are
amphipathic: the hydrophilic parts are
shown in blue, and the hydrophobic parts
are shown in black.

o ch 2 — o— c — ft

II I

R 2 — c— O — C— H O

ch 2 — o— c — r 3

Triacylglycerol

Pancreatic

lipase

2 H 2 0

0 O — C — R,
+

0 o— c— r 3

2 H®

O CH 2 — OH

II I

B. Lipoproteins

Triacylglycerols, cholesterol, and cholesteryl esters cannot be transported in blood or
lymph as free molecules because they are insoluble in water. Instead, these lipids assem-
ble with phospholipids and amphipathic lipid-binding proteins to form spherical

◄ lUMBM-Nicholson metabolic chart for lipid metabolism in mammals.

CH 2 — OH

2-Monoacylglycerol
▲ Figure 16.29

Action of pancreatic lipase. Removal of the
C-l and C-3 acyl chains produces free fatty
acids and a 2-monoacylglycerol. The inter-
mediates, 1,2- and 2 ,3-d iacylglycerol , are
not shown.

506 CHAPTER 16 Lipid Metabolism

Figure 16.30 ►

Action of phospholipase A 2 . X represents a
polar head group. Ri and R 2 are long hy-
drophobic chains, making up much of the
phospholipid molecule.

▲ Figure 16.31

Structure of phospholipase A 2 from cobra
venom. Phospholipase A 2 catalyzes the
hydrolysis of phospholipids at lipid-water
interfaces. The model shows how a
phospholipid substrate (dimyristoyl
phosphatidylethanolamine, space-filling
model) can fit into the active site of the
water-soluble enzyme. A calcium ion
(purple) in the active site probably helps
bind the anionic head group. About half of
the hydrophobic portion of the lipid would
be buried in the lipid aggregate. Mammalian
phospholipases are structurally similar to
the venom enzyme. [PDB 1POB].

1 G

0=P — 0°

1 2 I

H 2 C — CH— 3 CH 2

0 O

1 I

0= c c = o

I I

Ri R 2

Glycerophospholipid

1 0

0=P — 0°

Phospholipase A 2

H 2 0 O 0 +

C = 0

r 2

1 2 3 I

H 2 c— CH — CH 2

0 OH

o=c

R 1

Lysophosphoglyceride

macromolecular particles known as lipoproteins. A lipoprotein has a hydrophobic core
containing triacylglycerols and cholesteryl esters and a hydrophilic surface consisting of
a layer of amphipathic molecules such as cholesterol, phospholipids, and proteins
(Figure 16.32).

The largest lipoproteins are chylomicrons that deliver triacylglycerols and choles-
terol from the intestine via the lymph and blood to tissues such as muscle (for oxida-
tion) and adipose tissue (for storage) (Figure 16.33). Chylomicrons are present in blood
only after a meal. The cholesterol- rich remnants of chylomicrons — having lost most of
their triacylglycerol — deliver cholesterol to the liver. Liver cells are responsible for syn-
thesizing most of the newly synthesized cholesterol that enters the bloodstream but al-
most all cell types make cholesterol for internal use. Lipoproteins deliver both dietary
and liver-derived cholesterol to the rest of the body’s cells. Cholesterol biosynthesis is
regulated by hormones and by the levels of cholesterol in the blood.

Blood plasma contains several other types of lipoproteins. They are classified ac-
cording to their relative densities and types of lipid (Table 16.1). Since proteins are
more dense than lipids, the greater the protein content of a lipoprotein, the greater its
density. Very low density lipoproteins (VLDLs) consist of approximately 98% lipid and
only 2% protein. VLDLs are formed in the liver and carry lipids synthesized in the liver,
or not needed by the liver, to other tissues such as adipose tissue. Lipases within capil-
laries of muscle and adipose tissue degrade VLDLs and chylomicrons. When VLDLs
give up triacylglycerols to tissue cells their lipid content decreases and their remnants

BOX 16.6 EXTRA VIRGIN OLIVE OIL

Olive oil contains mostly triacylglycerols. If it has been pro-
duced by crushing olives with no additional chemical treat-
ment, then it is called virgin olive oil according to the Inter-
national Olive Oil Council (IOOC).

The quality of olive oil is often determined by the pres-
ence of free fatty acids that form when triacylglycerols break
down during production. Virgin olive oil should have less
than 2% free fatty acids (acidity) and extra virgin olive oil has
less than 0.8% free fatty acids (acidity).

► Extra virgin olive oil. Extra virgin olive oil has less than 0.8% free
fatty acids, http://www.examiner.com/fountain-of-youth-in-atlanta/
extra-virgin-olive-oil-benefits

16.10 Absorption and Mobilization of Fuel Lipids 507

are degraded to intermediate density lipoproteins (IDLs). Of the IDLs formed during
the breakdown of VLDLs, some are taken up by the liver and others are degraded to low
density lipoproteins (LDLs). LDLs are enriched in cholesterol and cholesteryl esters and
deliver these lipids to peripheral tissues. High density lipoproteins (HDLs) are formed
as protein-rich particles in blood plasma. They pick up cholesterol from peripheral tis-
sues, chylomicrons, and VLDL remnants and convert it into cholesterol esters. HDLs
transport cholesterol and cholesteryl esters back to the liver. Cholesteryl esters from
HDLs can be picked up by IDLs, which become LDLs.

Large lipoprotein particles contain a number of different lipid binding proteins.
These are often called apolipoproteins — the cc apo-” prefix usually refers to polypeptides
that bind to a tightly associated cofactor as described in Chapter 7. Two of these
apolipoproteins are large, hydrophobic, monomeric proteins. ApoB-100 (M r 513,000) is
firmly bound to the outer layer of VLDLs, IDLs, and LDLs. The smaller apolipoproteins
of VLDLs and IDLs are weakly bound and most dissociate during lipoprotein degrada-
tion, leaving apoB-100 as the major protein component of LDLs. ApoB-48 (M r
241,000), which is present only in chylomicrons, is identical in primary structure to the
N-terminal 48% of apoB-100.

ApoB-100 and apoB-48 form much of the amphipathic crust or shell over the hy-
drophobic lipoprotein core of their respective lipoproteins. ApoB-100 is the protein that
attaches LDL to its cell surface receptor; apoB-48 lacks this property. The other
apolipoproteins are smaller than apoB-48. They have a variety of functions, including
modulating the activity of certain enzymes involved in lipid mobilization and interact-
ing with cell surface receptors.

Cholesterol, an essential component of eukaryotic cell membranes, is delivered to
peripheral tissues by LDLs. The lipoprotein particles bind to the LDL receptor on the cell
surface. A complex between LDL and its receptor enters the cell by endocytosis and fuses
with a lysosome. Lysosomal lipases and proteases degrade the LDL releasing cholesterol
that is then incorporated into cell membranes or stored as cholesteryl esters.
An abundance of intracellular cholesterol suppresses synthesis of HMG-CoA reductase,
a key enzyme in the biosynthesis of cholesterol and it also inhibits synthesis of the LDL
receptor. Individuals lacking LDL receptors suffer from familial hypercholesterolemia, a
disease in which cholesterol accumulates in the blood and is deposited in the skin and
in arteries. Such patients die of heart disease at an early age.

HDLs remove cholesterol from plasma and from cells of nonhepatic tissues return-
ing it to the liver. They bind to a receptor called SR-B1 at the liver surface and transfer
cholesterol and cholesterol esters into liver cells. The lipid depleted HDL particles re-
turn to the plasma. In the liver, the cholesterol can be converted to bile salts that are se-
creted into the gallbladder.

The buildup of lipid deposits in the arteries (atherosclerosis) is associated with in-
creased risk of coronary heart disease that can lead to a heart attack. High levels of LDL
(“bad” cholesterol) increase the chance of developing atherosclerosis. High levels of

Core containing
triacylglycerols
and cholesteryl
esters

Cholesterol

Lipoprotein

▲ Figure 16.32

Structure of a lipoprotein. A core of neutral
lipids, including triacylglycerols and choles-
teryl esters, is coated with phospholipids in
which apolipoproteins and cholesterol are
embedded.

▲ Chylomicrons.

BOX 16.7 LIPOPROTEIN LIPASE AND CORONARY HEART DISEASE

Lipoprotein lipase (Section 16.9) is the enzyme that releases
fatty acid from the triacylglcerols in lipoproteins. It plays an
important role in clearing triacylglycerols from the blood
plasma. High concentrations of triacylglycerols are associ-
ated with coronary heart disease.

The human population contains several variants (muta-
tions) of the lipoprotein lipase (LPL) gene. Some of these are
associated with decreased LPL activity. One example is the
D9N variant where an asparagine residue substitutes for the
normal aspartate residue at position 9. Individuals who carry
this variant are more likely to suffer from coronary heart dis-
ease due to the buildup of triacyglycerol- containing lipopro-
teins in the blood plasma.

In the S447X variant a normal serine codon is mutated
to a stop codon (X) at position 447. The result is a truncated
protein that is shorter than the normal protein. About 17%
of the population carries at least one copy of this variant
gene and 1% of the population is homozygous for this vari-
ant. The S447X enzyme is more active than the wild-type
enzyme and this results in lower triacylglycerol levels in
plasma. Males (but not females) who carry this variant are
less likely to suffer heart attacks. This is an example of a ben-
eficial allele that has arisen in the human population.

[Online Mendelian Inheritance in Man (OMIM) MIM=609708]

508 CHAPTER 16 Lipid Metabolism

Figure 16.33 ►

Summary of lipoprotein metabolism.

Chylomicrons formed in intestinal cells carry
dietary triacylglycerols to peripheral tissues,
including muscle and adipose tissue. Chy-
lomicron remnants deliver cholesteryl esters
to the liver. VLDLs assemble in the liver and
carry endogenous lipids to peripheral tis-
sues. When VLDLs are degraded (via I DLs),
they pick up cholesterol and cholesteryl es-
ters from HDLs and become LDLs, which
carry cholesterol to nonhepatic tissues.

HDLs deliver cholesterol from peripheral
tissues to the liver.

INTESTINE LIVER

▲ Figure 16.34

Human serum albumin. Seven bound mole-
cules of palmitate are shown. [PDB 1E7H]

HDL (“good” cholesterol), on the other hand, are correlated with a decrease in the risk
of having a heart attack. Statins (Box 16.4) block synthesis of cholesterol in the liver and
lower LDL levels.

C. Serum Albumin

In addition to complex lipids such as cholesterol and triacylglycerols, free fatty acids are
also transported in blood plasma. Fatty acids bind to serum albumin, an abundant
plasma protein. This protein, especially the bovine version (bovine serum albumin,
BSA) has been intensely studied for over 40 years. Recently, the structure of human
serum albumin (HSA) in association with free fatty acids of various chain lengths
(Figure 16.34) has been solved by X-ray crystallography.

HSA belongs to the all -a category of tertiary structures (Section 4.7, Figure 4.24a).
There are seven distinct binding sites for palmitic acid (16:0) and other medium and
long chain fatty acids. In most cases, the carboxylate end of the fatty acids interacts with
the side chains of basic amino acid residues and the methylene tails fit into hydrophobic
pockets that can accommodate chains of 10-18 carbons. HSA also binds many impor-
tant drugs that are only sparingly soluble in water.

16.11 Ketone Bodies Are Fuel Molecules

Most acetyl CoA produced in the liver from fatty acid oxidation is routed to the citric
acid cycle but some of it can follow an alternate pathway. During periods of fasting, gly-
colysis is decreased and the gluconeogenic pathway is active. Under these conditions the

Table 16.1 Lipoproteins in human plasma

Chylomicrons

VLDLs

IDLs

LDLs

HDLs

Molecular weight x 10 -6

>400

10-80

5-10

2.3

0.18-0.36

Density (g cm -3 )

<0.95

0.95-1.006

1.006-1.019

1.019-1.063

1.063-1.210

Chemical composition (%)

Protein

Triacylglycerol

Cholesterol

Phospholipid

16.11 Ketone Bodies Are Fuel Molecules 509

pool of oxaloacetate molecules becomes temporarily depleted. The amount of acetyl
Co A from enhanced /3 - oxidation exceeds the capacity of the citric acid cycle (recall that
oxaloacetate reacts with acetyl CoA in the first step of the citric acid cycle). The excess
acetyl CoA is used to form ketone bodies — /3-hydroxybutyrate, acetoacetate, and ace-
tone. As indicated by their structures (Figure 16.35), not all ketone bodies are ketones.
The only quantitatively significant ketone bodies are /3-hydroxybutyrate and acetoac-
etate; small amounts of acetone are produced by the nonenzymatic decarboxylation of
acetoacetate, a /3-keto acid.

/3-Hydroxybutyrate and acetoacetate are fuel molecules. They have less potential
metabolic energy than the fatty acids from which they are derived but they make up for
this deficiency by serving as “water-soluble lipids” that can be more readily transported
in the blood plasma. During starvation, ketone bodies are produced in large amounts
becoming substitutes for glucose as the principal fuel for brain cells. Ketone bodies are
also metabolized in skeletal muscle and in the intestine during starvation.

A. Ketone Bodies Are Synthesized in the Liver

In mammals, ketone bodies are synthesized in the liver and exported for use by other
tissues. The pathway for ketone body synthesis is shown in Figure 16.36. First, two mol-
ecules of acetyl CoA condense to form acetoacetyl CoA and HS-CoA in a reaction cat-
alyzed by acetoacetyl- CoA thiolase. Subsequently, a third molecule of acetyl CoA is
added to acetoacetyl CoA to form 3-hydroxy-3-methylglutaryl CoA (HMG CoA) in a
reaction catalyzed by HMG-CoA synthase. These steps are identical to the first two steps
in the isopentenyl diphosphate biosynthesis pathway (Figure 16.17). The synthesis of

2 Acetyl CoA

HS-CoA

Acetoacetyl-CoA

thiolase

Acetoacetyl CoA

H 2 0 +

Acetyl CoA^\

HMG-CoA

HS-CoA«y s v nthase
+ H ® -

OH O

O 1 11

u ooc— ch 2 — c — ch 2 — C— S-CoA

ch 3

3-Hydroxy-3-methylglutaryl CoA (HMG CoA)

HMG-CoA

lyase

H 3 C—C— S-CoA
Acetyl CoA

G OOC — CH 2 — C — CH 3
Acetoacetate

NADH + H®

NAD

( 3 - Hyd roxy b uty rate
dehydrogenase

O 1

u ooc— ch 2 — c — ch 3

nonenzymatic

^ co 2

N /

h 3 c — c — ch 3

Acetone

/3-Hydroxybutyrate

0 1

u ooc— ch 2 — c — ch 3

/3-Hydroxybutyrate

u ooc— ch 2 — c — ch 3

Acetoacetate

h 3 c — c — ch 3

Acetone

▲ Figure 16.35
Ketone bodies.

◄ Figure 16.36

Biosynthesis of /?-hydroxybutyrate,
acetoacetate, and acetone.

510 CHAPTER 16 Lipid Metabolism

▲ HMG-CoA synthase. The human ( Homo
sapiens ) isozymes are shown with bound
HMG CoA. The cytosolic enzyme (top: PDB
2P8U) and the mitochondrial version
(bottom: PDB 2WYA) are very similar.

Changes in carbohydrate metabolism
during starvation are described in
Section 13.10.

ketone bodies takes place in mitochondria but the synthesis of isopentenyl diphosphate
(and cholesterol) takes place in the cytosol. Mammals have distinct isozymes of
acetoacetyl-CoA thiolase and HMG-CoA synthase in the mitochondria and the cytosol.
HMG-CoA synthase is only present in the mitochondria of liver cells and not in the mi-
tochondria of any other cell types.

In the next step, HMG-CoA lyase catalyzes the cleavage of HMG CoA producing
acetoacetate and acetyl CoA. HMG-CoA lyase is not present in the cytosol, which is why
cytosolic HMG CoA is used exclusively in isopentenyl diphosphate synthesis and no ke-
tone bodies are produced in the cytosol. NADH-dependent reduction of acetoacetate
produces /3-hydroxybutyrate in a reaction catalyzed by /3-hydroxybutyrate dehydroge-
nase. Both acetoacetate and /3-hydroxybutyrate can be transported across the inner mi-
tochondrial membrane and the plasma membrane of liver cells. They enter the blood to
be used as fuel by other cells of the body. Small amounts of acetoacetate are nonenzy-
matically decarboxylated to acetone in the bloodstream.

The main control point for ketogenesis is the mitochondrial isozyme of HMG-CoA
synthase provided that fatty acyl CoA and acetyl CoA are available in the mitochondria.
Succinyl CoA specifically inhibits this enzyme by covalent modification through suc-
cinylation. This is a short-term inactivation since reactivation occurs frequently by
spontaneous desuccinylation. Glucagon lowers the amount of succinyl CoA in mito-
chondria, stimulating ketogenesis. Long-term regulation occurs by modification of
gene expression. Starvation increases the level of HMG-CoA synthase (and its mRNA);
refeeding or insulin produces a decrease in both activity and mRNA.

B. Ketone Bodies Are Oxidized in Mitochondria

In cells that use them as an energy source, /3-hydroxybutyrate and acetoacetate enter
mitochondria where they are converted to acetyl CoA that is oxidized by the citric acid
cycle. /3-Hydroxybutyrate is converted to acetoacetate in a reaction catalyzed by an
isozyme of /3-hydroxybutyrate dehydrogenase that is distinct from the liver enzyme.
Acetoacetate reacts with succinyl CoA to form acetoacetyl CoA in a reaction catalyzed
by succinyl-CoA transferase (also called succinyl-CoA:3-ketoacid-CoA transferase;
Figure 16.37). Ketone bodies are broken down only in nonhepatic tissues because this
transferase is present in all tissues except liver. The succinyl- CoA transferase reaction
siphons some succinyl CoA from the citric acid cycle. Energy that would normally be
captured as GTP in the substrate-level phosphorylation catalyzed by succinyl- CoA syn-
thetase (Section 13.3#5) is used instead to activate acetoacetate to its CoA ester. Thiolase
then catalyzes the conversion of acetoacetyl CoA to two molecules of acetyl CoA that
can be oxidized by the citric acid cycle.

O O

HS-CoA-

Thiolase

Figure 16.37 ►

Conversion of acetoacetate to acetyl CoA.

H 3 C — C— S-CoA
Acetyl CoA

Problems 511

BOX 16.8 LIPID METABOLISM IN DIABETES

The breakdown of fats occurs because lipolysis is not inhib-
ited by insulin, and other hormones trigger the release of
fatty acids from adipocytes. The large amounts of fatty acids
available to the liver lead to excess acetyl CoA that is diverted
to form ketone bodies. In Type 2 diabetes (Section 12.7), the
accumulation of glucose in the blood is caused mainly by
poor uptake of glucose by peripheral tissues. Because obesity
strongly predisposes a person to developing Type 2 diabetes,
much research is focusing on the role of lipids in decreased
insulin sensitivity. It appears that elevated free fatty acids in
the blood may interfere with insulin signaling for glucose up-
take into tissues.

Individuals suffering from untreated Type 1 diabetes
produce large amounts of ketone bodies — more than the
peripheral tissues can use. The smell of acetone can be dis-
cerned on the breath of diabetics. In fact, the levels of ace-
toacetic acid and /3-hydroxybutyric acid in the blood can be
so high that the pH of the serum can be lowered — a life-
threatening condition called diabetic ketoacidosis. Type 1

diabetes must be treated with repeated injections of insulin
and restricted glucose intake.

Although acute complications are rare in Type 2 dia-
betes, hyperglycemia can lead to tissue damage, particularly
in the eye and the cardiovascular and renal systems. Dietary
modifications are often sufficient to control Type 2 diabetes.
In addition, oral drugs can increase insulin secretion and po-
tentiate its action at peripheral tissues.

A novel approach for the treatment of Type 2 diabetes
may be inhibition of the tyrosine phosphatase PTP-1B. PTP-
1B inactivates the insulin receptor by catalyzing the removal
of phosphate added to the receptor when insulin binds to it.
After insulin injection, mice lacking PTP-1B have increased
phosphorylation of insulin receptors in liver and muscle and
enhanced sensitivity to insulin. These mice also maintain
normal levels of blood glucose after a meal. A surprising ob-
servation was that mice lacking PTP-1B could eat a high fat
diet yet be resistant to weight gain. PTP-1B may therefore
also be a target for the treatment of obesity.

Summary

1. The pathway for fatty acid synthesis begins with synthesis of mal-
onyl CoA in a reaction catalyzed by acetyl CoA- carboxylase. Mal-
onyl CoA is converted to malonyl ACP and one molecule of
malonyl ACP condenses with acetyl CoA (or acetyl ACP) to form
acetoacetyl ACP.

2. The formation of long- chain fatty acids from a 3-ketoacyl ACP
precursor occurs in four stages: reduction, dehydration, further
reduction, and condensation . These four stages repeat to form a
long-chain fatty acid. Fatty acids with more than 18 carbons and
unsaturated fatty acids are produced by additional reactions.

3. Triacylglycerols and glycerophospholipids are derived from
phosphatidate. The synthesis of triacylglycerols and neutral
phospholipids proceeds via a 1,2-diacylglycerol intermediate.
Acidic phospholipids are synthesized via a CDP-diacylglycerol
intermediate.

4. Many eicosanoids are derived from arachidonate. The cyclooxy-
genase pathway leads to prostacyclin, prostaglandins, and throm-
boxane A 2 . The products of the lipoxygenase pathway include
leukotrienes.

5. Sphingolipids are synthesized from serine and palmitoyl CoA. Re-
duction, acylation, and oxidation produce ceramide, which can be
modified by adding a polar head group and sugar residues.

6. Cholesterol is synthesized from acetyl CoA in a pathway leading
to mevalonate and isopentenyl diphosphate. Both cholesterol

and isopentenyl diphosphate are precursors of many other
compounds.

7. Fatty acids are degraded to acetyl CoA by /3-oxidation, the se-
quential removal of two-carbon fragments. Fatty acids are first
activated by esterification to CoA and fatty acyl CoA is oxidized
by a repeated series of four enzyme- catalyzed steps: oxidation,
hydration, further oxidation, and thiolysis. Fatty acids yield more
ATP per gram than glucose.

8. /3-Oxidation of odd-chain fatty acids produces acetyl CoA and
one molecule of propionyl CoA. The oxidation of most unsatu-
rated fatty acids requires two enzymes, an isomerase and a reduc-
tase, in addition to those required for the oxidation of saturated
fatty acids.

9. Fatty acid oxidation in animals is regulated by hormones accord-
ing to the energy needs of the organism.

10 . Dietary fat is hydrolyzed in the intestine to fatty acids and
monoacylglycerols, which are absorbed. Lipoproteins transport
lipids in the blood. In adipocytes, fatty acids are esterified for
storage as triacylglycerols. Fatty acids are mobilized by the action
of hormone-sensitive lipase.

11. The ketone bodies /3-hydroxybutyrate and acetoacetate are water-
soluble fuel molecules produced in the liver by the condensation
of acetyl- CoA molecules.

Problems

1. (a) Familial hypercholesterolemia is a human genetic disorder in
which LDL receptors are defective, leading to very high blood
cholesterol levels and severe atherosclerosis at an early age.
Explain why this disease results in high blood cholesterol
levels.

(b) Do high blood cholesterol levels affect cellular cholesterol
synthesis in individuals with this disease?

(c) Individuals with Tangier’s disease lack the cellular protein
ABC1, which is required for cholesterol uptake by HDL. How
will this disease affect cholesterol transport?

512 CHAPTER 16 Lipid Metabolism

2. Individuals with abnormally low levels of carnitine in their mus-
cles suffer from muscular weakness during moderate exercise. In
addition, their muscles have significantly increased levels of tria-
cylglycerols.

(a) Explain these two effects.

(b) Can these individuals metabolize muscle glycogen aerobi-
cally?

3. How many ATP equivalents are generated by the complete oxida-
tion of (a) laurate (dodecanoate) and (b) palmitoleate (cis- A 9 -
hexadecenoate)? Assume that the citric acid cycle is functioning.

4 . Tetrahydrolipstatin (Orlistat) is a drug treatment for obesity. It is
an inhibitor of pancreatic lipase. Suggest a rationale for use of
tetrahydrolipstatin to treat obesity.

5. In addition to the enzymes of /3-oxidation, what enzymes are nec-
essary to degrade the following fatty acids to acetyl CoA or acetyl
CoA and succinyl CoA?

(a) oleate (o'sCH 3 (CH 2 ) 7 CH = CH(CH 2 ) 7 COO e )

(b) arachidonate

(all cis CH 3 (CH 2 ) 4 (CH = CHCH 2 ) 4 (CH 2 ) 2 COO q )

6. Animals cannot carry out a net conversion of even chain fatty
acid carbons to glucose. On the other hand, some of the carbons
in odd- chain fatty acids can be gluconeogenic precursors to glu-
cose. Explain.

7. Where is the labeled carbon found when the following molecules
are added to a liver homogenate carrying out palmitate synthesis?

(a) H 14 C0 3 ®

(b) II

H 3 14 C— c— s— CoA

8. Triclosan (2,4,4-trichloro-2-hydroxydiphenyl ether) is an effec-
tive antimicrobial agent that is used in a wide range of consumer
products including soaps, toothpaste, toys, and cutting boards.
Triclosan is effective against a broad spectrum of bacteria and
mycobacteria and is an inhibitor of type II FAS enoyl acyl carrier
protein reductase.

(a) What reaction is catalyzed by enoyl acyl carrier protein re-
ductase?

(b) Why is enoyl acyl carrier protein reductase an appropriate
target for antimicrobials?

9. It has been proposed that malonyl CoA may be one of the signals
sent to the brain to decrease the appetite response. When mice are
given a derivative of cerulenin (a fungal epoxide) named C75,
their appetite is suppressed and they rapidly lose weight. Ceru-
lenin and its derivatives have been shown to be potent inhibitors
of fatty acid synthase (FAS). Suggest how C75 might act as a po-
tential weight reduction drug.

10 . (a) Draw a general pathway for converting carbohydrates to fatty

acids in a liver cell, and indicate which processes occur in the
cytosol and which occur in motochondria.

(b) About half the reducing equivalents necessary for fatty acid
synthesis are generated by glycolysis. Explain how these re-
ducing equivalents can be used for fatty acid synthesis.

11. (a) Acetyl CoA carboxylase (ACC), a key regulator for fatty acid

synthesis, exists in two different interconvertible forms:

(1) an active filamentous polymer (dephosphorylated), and

(2) an inactive protomeric form (phosphorylated). Citrate
and palmitoyl CoA can regulate fatty acid synthesis by prefer-
entially binding tightly to and stabilizing different forms of
ACC. Explain how each of these regulator functions by inter-
acting with ACC.

Filamentous polymer (active) ^ Protomer (inactive)

(b) What role do glucagon and epinephrine play in regulating
fatty acid synthesis?

12. Obesity is a serious health problem worldwide due in part to in-
creased food intake and reduced physical activity. Obesity is asso-
ciated with a variety of human disease including Type 2 diabetes
and cardiovascular diseases. Selective and specific inhibitors of
acetyl-CoA carboxylase have been proposed as potential anti-
obesity drugs.

(a) What effect would an inhibitor of acetyl-CoA carboxylase
have on fatty acid synthesis and fatty acid oxidation?

(b) One such inhibitor of acetyl-CoA carboxylase is CABI (struc-
ture below). What structural feature of CABI makes it a po-
tential acetyl-CoA carboxylase inhibitor? (Levert, K. L., Wal-
drop, G. L., Stephens, J. M. (2002). /. Biol Chem. A biotin
analog inhibits acetyl CoA carboxylase activity and adipoge-
nesis. 277:16347-16350.)

13. Write the equation for the conversion of eight acetyl CoA mole-
cules to palmitate.

14 . (a) In response to tissue damage in such injuries as heart attacks

and rheumatoid arthritis, inflammatory cells (e.g., mono-
cytes and neutrophils) invade the injured tissue and promote
the synthesis of arachadonic acid. Explain the reason for this
response.

(b) The biosynthesis of eicosanoids is affected by nonsteroidal
drugs such as aspirin and ibuprofen and by steroidal drugs such
as hydrocortisone and prednisone (which inhibit a specific
phospholipase). Why do steroidal drugs inhibit the biosynthesis
of both prostaglandins and leukotrienes, whereas aspirin-like
drugs inhibit the biosynthesis of only prostaglandins?

15 . Draw the correct structures of the following complex lipids.

(a) Phosphatidyl glycerol.

(b) Ethanolamine plasmalogen (l-alkyl-2-glycero-3-phospho-
ethanolamine).

16 . Excess dietary fat can be converted to cholesterol in the liver.
When palmitate labeled with 14 C at every odd-numbered carbon
is added to a liver homogenate, where does the label appear in
mevalonate?

17 . The therapeutic anti-inflammatory effects of aspirin arise from its
inhibition of the enzyme cyclooxygenase-2 (COX-2) — involved in
the synthesis of prostaglandins, mediators of inflammation, pain,
and fever. Aspirin irreversibly inhibits COX-2 by covalently

Selected Readings 513

transferring an acetyl group to a serine residue at the enzyme ac-
tive site. However, the undesirable side effect of stomach irritation
arises from the irreversible inhibition of the related intestinal en-
zyme cyclooxygenase- 1 (COX-1) by aspirin. COX-1 is involved in
the synthesis of prostaglandins that regulate secretion of gastric
mucin, which protects the stomach from acid. The aspirin analog
APHS was synthesized and shown to be 60 times more selective as
an inhibitor of COX-2 than of COX-1, suggesting that it could be
an anti-inflammatory drug with far less gastrointestinal side
effects. Draw the structure of the inactivated COX-2 enzyme-
inhibitor complex with APHS. Since aspirin and structural analogs

act at the active site of COX enzymes, will they exhibit competitive
inhibition patterns?

APHS

Selected Readings

General

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Vance, J. E., and Vance, D. E., eds. (2008).
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Lipid Synthesis

Athenstaedt, K., and Daum, G. (1999). Phospha-
tidic acid, a key intermediate in lipid metabolism.
Eur. J. Biochem. 266:1-16.

Frye, L. L., and Leonard, D. A. (1999). Lanosterol
analogs: dual- action inhibitors of cholesterol
biosynthesis. Crit. Rev. Biochem. Mol. Biol. 34:123-124.

Kent, C. (1995). Eukaryotic phospholipid synthe-
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Leibundgut, M., Maier, T., Jenni, S., and Ban, N.
(2008). The multienzyme architecture of eukary-
otic fatty acid synthases. Curr. Opin. Struct. Biol.
18:714-726.

Simmons, D. L., Botting, R. M., and Hla, T. (2004).
Cyclooxygenase isozymes: the biology of
prostaglandin synthesis and inhibition. Pharmacol.
Rev. 56:387-437.

Sommerville, C., and Browse, J. (1996). Dissecting
desaturation: plants prove advantageous. Trends in
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Wallis, J. G., Watts, J. L., and Browse, J. (2002).
Polyunsaturated fatty acid synthesis: what will
they think of next? Trends Biochem. Sci.
27:467-473.

White, S. W. Zheng, J., Zhang, Y-M., and Rock,

C. O. (2005). The structural biology of type II
fatty acid biosynthesis. Ann. Rev. Biochem.
74:791-831.

Lipid Catabolism

Bartlett, K., and Eaton, S. (2004). Mitochondrial
/3- oxidation. Eur. J. Biochem. 271:462-469.

Candlish, J. (1981). Metabolic water and the
camel’s hump — a textbook survey. Biochem Ed.
9:96-97.

Ishikawa, M., Tsuchiya, D., Oyama, T., Tsunaka, D.,
and Morikawa, K. (2004). Structural basis for
channelling mechanism of a fatty acid (3 - oxidation
multienzyme complex. EMBO J. 23:2745-2754.

Kim, J-J., and Battaile, K. P. (2002). Burning fat:
the structural basis of fatty acid /3- oxidation. Curr.
Opin. Struct. Biol. 12:721-728.

Toogood. H. S., van Thiel, A., Basran, J., Sutcliffe,
M. J., Scrutton, N. S., and Leys, D. (2004). Extensive
domain motion and electron transfer in the
human electron transferring flavoprotein medium
chain acyl-CoA dehydrogenase complex. /. Biol.
Chem. 279: 32904-32912.

Toogood, H. S., van Thiel, A., Scrutton, N. S., and
Leys, D. (2005). Stabilization of non-productive
conformations underpins rapid electron transfer
to electron-transferring flavoprotein. /. Biol.
Chem. 280:30361-30366.

Wanders, R. J. A., and Waterman, H. R. (2006).
Biochemistry of mammalian peroxisomes revis-
ited. Ann. Rev. Biochem. 75: 295-332.

Lipoproteins

Bhattacharya, A. A., Grime, T., and Curry, S.
(2000). Crystallographic analysis reveals common
modes of binding of medium- and long-chain
fatty acids to human serum albumin. /. Mol. Biol.
303:721-732.

Fidge, N. H. (1999). High density lipoprotein re-
ceptors, binding proteins, and ligands. /. Lipid Res.
40:187-201.

Gagne, S. E., Larson, M. G., Pimstone, S. N.,
Schaefer, E. J., Kastelein, J. J. P., Wilson, P. W. F.,
Ordovas, J. M., and Hayden, M. R. (1999). A com-
mon truncation variant of lipoprotein lipase
(S447X) confers protection against coronary heart
disease: the Framingham Offspring Study. Clin.
Genet. 55:450-454.

Kreiger, M. (1998). The “best” of cholesterols, the
“worst” of cholesterols: a tale of two receptors.
Proc. Natl. Acad. Sci. USA 95:4077-4080.

o c

o o

Amino Acid Metabolism

A uthors writing a chapter on amino acid metabolism have a nearly impossible
task. Any description will be incomplete since there are 20 different amino
acids and many intermediates in the biosynthesis and degradation pathways.
Furthermore, alternate pathways are used by different tissues, organelles, and organ-
isms. Fortunately, metabolic highlights can show the biological rationale of how amino
acids are formed and degraded without getting into excessive detail. Here we describe a
number of these highlights in order to illustrate the principles and concepts of amino
acid metabolism.

The metabolism of amino acids includes hundreds of enzymatic interconversions
of small molecules. Many of these reactions involve nitrogen atoms. Some of the inter-
mediates appear in the metabolic pathways described in preceding chapters but many
are described here for the first time. Although amino acids from the degradation of pro-
teins can be a source of energy, we are more concerned with their biosynthesis. Life is
compromised if all the amino acids are not available at the same time for protein syn-
thesis. We can consider the metabolism of the 20 common amino acids from two points
of view: the origins and fates of their nitrogen atoms and the origins and fates of their
carbon skeletons.

The abilities of organisms to synthesize amino acids differ widely. A few organisms
can assimilate N 2 and simple carbon compounds into amino acids — in other words,
they are totally self-supporting for amino acid synthesis. Other species can synthesize
the carbon chains of amino acids but require nitrogen in the form of ammonia. We
begin this chapter with an overview of the principles of nitrogen metabolism.

Some species cannot synthesize the carbon skeletons of every amino acid. Mam-
mals, for example, can make only about half of the amino acids they require; the rest —
called essential amino acids — must be obtained from the diet. Nonessential amino acids
are those that mammals can synthesize in sufficient quantity, provided they receive ade-
quate total dietary protein.

The routes for disposal of the nitrogen- containing waste products of amino acid
metabolism also vary among species. For example, excess nitrogen is excreted by aquatic
animals as ammonia, by birds and most reptiles as uric acid, and by many other terrestrial

We live now in the "Age of Bacteria. "
Our planet has always been in the
"Age of Bacteria , " ever since the first
fossils — bacteria , of course — were
entombed in rocks more than 3 and
a half billion years ago. On any pos-
sible , reasonable , or fair criterion ,
bacteria are — and always have
been — the dominant forms of life
on Earth.

Stephen Jay Gould, (1996),
Full House , p. 1 76

Top: Glutamine synthetase from the bacterium Salmonella typhimurium. Twelve identical subunits are arranged with
hexagonal symmetry. [PDB 2GLS].

514

17.1 The Nitrogen Cycle and Nitrogen Fixation 515

vertebrates as urea. We will end the chapter with a description of the urea cycle, a path-
way for elimination of nitrogen in mammals.

17.1 The Nitrogen Cycle and Nitrogen Fixation

The nitrogen needed for amino acids (and for the heterocyclic bases of nucleotides;
Chapter 18) comes from two major sources — nitrogen gas in the atmosphere and ni-
trate (N0 3 ®) in soil and water. Atmospheric N 2 , which constitutes about 80% of the at-
mosphere, is the ultimate source of biological nitrogen. This molecule can be metabo-
lized, or fixed, by only a few species of bacteria. N 2 and N0 3 ® must be reduced to
ammonia in order to be used in metabolism. The ammonia produced is incorporated
into amino acids via glutamate, glutamine, and carbamoyl phosphate.

N 2 is chemically unreactive because of the great strength of the N = N triple bond.
Some bacteria have a very specific, sophisticated enzyme, called nitrogenase, that can
catalyze the reduction of N 2 to ammonia in a process called nitrogen fixation. Ammonia
is essential for life and bacteria are the only organisms capable of producing it from at-
mospheric nitrogen. Half of all biological nitrogen fixation is performed by various
species of cyanobacteria in the ocean. The other half comes from soil bacteria.

There are two additional nitrogen-converting processes in addition to biological
nitrogen fixation. During lightning storms, high-voltage discharges catalyze the oxida-
tion of N 2 to nitrate and nitrite (N 0 2 ®). Nitrogen is converted to ammonia for use in
plant fertilizers by an energetically expensive industrial process that requires high tem-
perature and pressure as well as special catalysts to drive the reduction of N 2 by H 2 . The
availability of biologically useful nitrogen is often a limiting factor for plant growth, and
the application of nitrogenous fertilizers is important for obtaining high crop yields.
Humans are now responsible for a substantial fraction of the total nitrogen fixation on
the planet. Although only a small percentage of the nitrogen undergoing metabolism
comes directly from nitrogen fixation, this process is the only way that organisms can
use the huge pool of atmospheric N 2 .

The overall scheme for the interconversion of the major nitrogen-containing com-
pounds is shown in Figure 17.1. The flow of nitrogen from N 2 to nitrogen oxides, am-
monia, and nitrogenous biomolecules and then back to N 2 is called the nitrogen cycle.
Most of the nitrogen shuttles between ammonia and nitrate. Ammonia from decayed
organisms is oxidized by soil bacteria to nitrate. This formation of nitrate is called nitri-
fication. Some anaerobic bacteria can reduce nitrate or nitrite to N 2 (denitrification).

Metabolic _
pathways *

Amino acids
Nucleotides
Phospholipids

▲ Figure 17.1

Nitrogen cycle. A few free-living or symbiotic microorganisms can convert N 2 directly to ammonia.
Ammonia is incorporated into biomolecules such as amino acids and proteins that subsequently are
degraded, re-forming ammonia. Many soil bacteria and plants can carry out the reduction of nitrate
to ammonia via nitrite. Several bacteria convert ammonia to nitrite. Others oxidize nitrite to nitrate
and some can reduce nitrate to N 2 .

▲ Blooms of Trichodesmium. Trichodesmium
is one of the main nitrogen-fixing species of
cyanobacteria. This large bloom of bacteria
formed giant streaks in the ocean off the
coast of Australia. The photograph was
taken from the space shuttle. The average
concentration of nitrogen-fixing bacteria in
the ocean is about one million cells per liter.

KEY CONCEPT

Nitrogen is the most abundant gas in the
atmosphere but only a few species of
bacteria are capable of nitrogen fixation.

▲ Lightning. Lightning causes the conversion
of nitrogen gas to nitrates. It is an important
source of usable nitrogen for living organisms.
This photograph was taken in 1908.

516 CHAPTER 17 Amino Acid Metabolism

▲ Figure 17.2

Nodules on alfalfa roots. Symbiotic bacteria
of the genus Rhizobium reside in these nod-
ules where they reduce atmospheric nitrogen
to ammonia.

Most green plants and some microorganisms contain nitrate reductase and nitrite re-
ductase, enzymes that together catalyze the reduction of nitrogen oxides to ammonia.

2e 0 # 2H @

FLO

6e° f 7H @ 2 H 2 0

Nitrate

■ no 2 ° ■

Nitrite

NH 3

Ammonia

(17.1)

This ammonia is used by plants, which supply amino acids to animals. Reduced ferre-
doxin (formed in the light reactions of photosynthesis, Section 15.2B) is the source of
the reducing power in plants and photosynthetic bacteria.

Let s examine the enzymatic reduction of N 2 . Most nitrogen fixation in the biosphere
is carried out by bacteria that synthesize the enzyme nitrogenase. This multisubunit pro-
tein catalyzes the conversion of each molecule of N 2 to two molecules of NH 3 . Nitroge-
nase is present in various species of Rhizobium and Bradyrhizobium that live symbiotically
in root nodules of many leguminous plants, including soybeans, peas, alfalfa, and clover
(Figure 17.2). N 2 is also fixed by free-living soil bacteria such as Agrobacteria, Azotobacter,
Klebsiella, and Clostridium and by cyanobacteria (mostly Trichodesmuim spp .) found in
the ocean. Most plants require a supply of fixed nitrogen from sources such as decayed an-
imal and plant tissue, nitrogen compounds excreted by bacteria, and fertilizers. Verte-
brates obtain fixed nitrogen by ingesting plant and animal matter.

Nitrogenase is a protein complex that consists of two different polypeptide sub-
units forming an a 2 l 3 2 dimer of dimers (Figure 17.3). The two halves of the complex
contain an [8 Fe-7 S] iron-sulfur cluster called the P- cluster. It is near the outer surface
of the protein. The reactive center is a complex cluster of molybdenum, iron, and ho-
mocitrate [MoFe 7 S 9 -homocitrate]. A single a/3 dimer is called the iron-molybdenum
(MoFe) protein.

Electrons are donated to the P-custer by a mobile iron (Fe) protein containing a
[4 Fe-4 S] cluster. Fe protein, a homodimer, binds to the ends of MoFe protein near the
P- cluster and a single electron is passed from Fe protein to MoFe protein. The reduction
of iron in Fe protein is coupled to oxidation of ferredoxin or flavodoxin and each of these
reduction reactions requires hydrolysis of two bound ATP molecules. Electrons are
passed from Fe protein to the P-cluster to the FeMo-cluster. A total of six electrons are re-
quired for conversion of N 2 to 2NH 3 and these must be passed one at a time from Fe
protein as it binds and then dissociates from MoFe protein. An obligatory reduction of 2
H® to H 2 accompanies the reduction of N 2 . The overall stoichiometry is

N 2 + 8 H© + 8 e 0 + 1 6 ATP — » 2 NH 3 + H 2 + 1 6 ADP + 1 6 P, (17.2)

Figure 17.3 ►

Structure of Azotobacter vinelandii
nitrogenase. The Fe protein subunits are
colored red and orange and the a and p sub-
units of each half of the MoFe protein are
colored blue/green and purple/pink. This
structure with bound Fe protein is stabilized
by bound transition-state ATP analogs ADP-
AIF 4 at the ATP binding sites. [PDB 1N2C]

Fe Protein MoFe Protein MoFe Protein Fe Protein

n r

[4 Fe-4 S] cluster

P-cluster

MoFe 7 S 9 N-homocitrate
reactive center

17.1 The Nitrogen Cycle and Nitrogen Fixation

517

This is a very expensive reaction in terms of ATP equivalents. It is also a very slow reac-
tion in biochemical terms with a turnover number of only five ammonia atoms pro-
duced per second. The slowness of the reaction is due to the fact that eight reduced Fe
proteins have to bind and dissociate from the MoFe protein during the conversion of
nitrogen to ammonia.

Nitrogenases must be protected from oxygen because the various oxidation-reduction
centers are highly susceptible to inactivation by 0 2 . Strict anaerobes carry out nitrogen fixa-
tion in the absence of 0 2 . Within the root nodules of leguminous plants, the protein leghe-
moglobin (a homolog of vertebrate myoglobin; Section 4. 12) binds 0 2 and thereby keeps its
concentration sufficiently low in the immediate environment of the nitrogen- fixing
enzymes of Rhizobium. Nitrogen fixation in cyanobacteria is carried out in specialized cells
(heterocysts) whose thick membranes inhibit entry of 0 2 (Figure 10.8). In order to obtain
the reducing power and ATP required for this process, symbiotic nitrogen-fixing microor-
ganisms rely on nutrients obtained through photosynthesis carried out by the plants with
which they are associated.

The actual reduction of nitrogen takes place at the iron-molybdenum-homocitrate
cluster in the MoFe protein. This cluster is remarkably complex. It consists of a cage of
Fe and S atoms surrounding a central N atom. A single Mo atom is bound to one edge
of the Fe — S cage. It is chelated to a single molecule of homocitrate to form a
MoFe 7 S 9 N*homocitrate cluster (Figure 17.4).

The detailed reaction mechanism of nitrogenase is unknown in spite of many years
of intense study. It is likely that each of the three N = N bonds is broken sequentially,
giving rise to the intermediates diimine and hydrazine.

2e e , 2H© 2e e ,2H© 2e e , 2H©

N = N > H — N = N — H » H 2 N — NH 2 » NH 3 + NH 3

Diimine Hydrazine (17.3)

The reduction of 2 H© to H 2 , an essential coupled reaction, consumes the extra
pair of electrons from ferredoxin as shown in Reaction 17.2.

(a)

O Carbon
O Hydrogen
Q Oxygen
03 Nitrogen

Q Sulfur
Q Iron

Q Molybdenum

◄ Figure 17.4

Structure of the MoFe 7 S 9 N • homocitrate reac-
tive center in Azotobacter vinelandii. (a) Rest-
ing state, (b) One possible structure with
bound N 2 . [PDB 2MIN]

518 CHAPTER 17 Amino Acid Metabolism

coo®

NAD(P)H + H®

coo®

COO®

„C=0

I NAD(P)®

HoN — C — H

ATP ADP+P;

1 T

H,N — CH

a |

ch 2 + nh 4 ®

v / +H ^°

a a |

cn 2

/Glutamine

Glutamate dehydrogenase

1 synthetase

NH®

0 0°

^ C \

a-Ketoglutarate

Glutamate

Glutamine

a Figure 17.5

Incorporation of ammonia into glutamate and glutamine.

Synthetases are members of the Ligase
class of enzymes. They require ATP as a
cosubstrate. Synthases are members of
the Transferase or Lyase class of
enzymes. They do not use ATP as a
cofactor. (Section 5.1, Section 13.3#1).

coo 0

H 3 N — C — H

Glutamine

coo 0

C =0

oh 2

^ \ (P)

O 0°

u-Ketoglutarate

Glutamate

synthase

NAD(P)H

+ H®

NAD(P)®

COO

,©

H 3 N — C — H

ChH,

ch 2

,©

2 Glutamate

▲ Figure 17.6

Glutamate synthase catalyzes the reductive
amination of a-ketoglutarate.

17.2 Assimilation of Ammonia

Ammonia is assimilated into a large number of low molecular weight metabolites, often
via the amino acids glutamate and glutamine. At physiological pH the main ionic form
of ammonia is the ammonium ion, NH 4 ® (piC a = 9.2). However, unprotonated ammonia
(NH 3 ) is the reactive species in the catalytic centers of many enzymes.

A. Ammonia Is Incorporated into Glutamate and Glutamine

The reductive amination of a-ketoglutarate to glutamate by glutamate dehydrogenase is
one highly efficient route for the incorporation of ammonia into the central pathways
of amino acid metabolism (Figure 17.5). The glutamate dehydrogenases of some species
or tissues are specific for NADH while others are specific for NADPH. Still others can
use either cofactor.

The glutamate dehydrogenase reaction can play different physiological roles de-
pending on substrate and coenzyme availability and enzyme specificity. In
Escherichia coli , for example, the enzyme generates glutamate when NH 4 ® is present
at high concentrations. In the mold Neurospora crassa an NADPH-dependent en-
zyme is used for the reductive amination of a-ketoglutarate to glutamate and the re-
verse reaction is catalyzed by an NAD® -dependent enzyme. Glutamate dehydroge-
nase is located in mitochondria in mammals and plants and it catalyzes a near
equilibrium reaction with net flux usually from glutamate to a-ketoglutarate. The
primary role of glutamate dehydrogenase in mammals is the degradation of amino
acids and the release of NH 4 ® . Mammals probably assimilate very little nitrogen as
free ammonia because they get most of their nitrogen from amino acids and nu-
cleotides in the diet.

Another reaction critical to the assimilation of ammonia in many organisms is the
formation of glutamine from glutamate and ammonia. This reaction is catalyzed by
glutamine synthetase (Figure 17.5). Glutamine is a nitrogen donor in many biosyn-
thetic reactions; for example, the amide nitrogen of glutamine is the direct precursor of
several of the nitrogen atoms of the purine and pyrimidine ring systems of nucleotides
(Sections 18.1 and 18.3). In mammals, glutamine carries nitrogen and carbon between
tissues in order to avoid high levels of toxic NH 4 ® in the bloodstream.

The amide nitrogen of glutamine can be transferred to a-ketoglutarate to produce
two molecules of glutamate in a reductive amination reaction catalyzed by glutamate
synthase (Figure 17.6). Like glutamate dehydrogenase, glutamate synthase requires a re-
duced pyridine nucleotide to reductively aminate a-ketoglutarate. Unlike the dehydro-
genase, the synthase uses glutamine as the source of nitrogen. Animals do not have glu-
tamate synthase.

B. Transamination Reactions

The amino group of glutamate can be transferred to many a-keto acids in reactions cat-
alyzed by enzymes known as transaminases or aminotransferases. The general transam-
ination reaction is shown in Figure 17.7.

17.2 Assimilation of Ammonia

519

▲ Pig ( Sus scrofa ) cytosolic aspartate transaminase. The enzyme is a
dimer of identical subunits (individual monomers are shown in purple
and blue). A molecule of the coenzyme pyridoxal phosphate is shown
(space-filling model) in each active site. [PDB 1 AJR]

The amino group of glutamate is transferred to various ct-keto acids generating the
corresponding a -amino acids during amino acid synthesis. Most of the common amino
acids can be formed by transamination. In amino acid catabolism, amino groups are
transferred from various amino acids to a-ketoglutarate or oxaloacetate generating glu-
tamate or aspartate.

All known transaminases require the coenzyme pyridoxal phosphate (Section 7.8).
The chemical mechanism of the initial half- reaction of transamination was shown in
Figure 7.18. The complete transamination requires two coupled half-reactions, with
enzyme-bound pyridoxamine phosphate (PMP) transiently carrying the amino group
being transferred.

The transaminases catalyze near- equilibrium reactions. The direction in which the re-
actions proceed in vivo (flux) depends on the supply of substrates and the removal of prod-
ucts. For example, in cells with an excess of a- amino nitrogen groups the amino groups
can be transferred via one or a series of transamination reactions to a-ketoglutarate to
yield glutamate that can undergo oxidative deamination catalyzed by glutamate dehydro-
genase. Transamination occurs in the opposite direction when amino acids are being ac-
tively formed and the amino groups are donated by glutamate.

An important alternative to the glutamate dehydrogenase reaction in bacteria uses
coupled reactions catalyzed by glutamine synthetase and glutamate synthase for the as-
similation of ammonia into glutamate, especially when the concentration of ammonia
is low. Figure 17.8 shows how the combined actions of glutamine synthetase and gluta-
mate synthase can lead to the incorporation of ammonia into a variety of amino acids.
After formation, glutamate undergoes transamination with ct-keto acids to form the
corresponding amino acids. The conversion of a-ketoglutarate to glutamate can occur
via the glutamine synthetase-glutamate synthase pathway at the low concentrations of
NH 4 ® present in most bacterial cells because the K m of glutamine synthetase for NH 3
is much lower than the K m of glutamate dehydrogenase for NH 4 ®.

coo°

H 3 N — CH

R i

(u-Amino acid)!

0 =

COO

■ c

R 1

(a-Keto acid).

COO°

0= c
I

R 2

(u-Keto acid) 2

H 3 N — CH

R 2

(u-Amino acid) 2

▲ Figure 17.7

Transfer of an amino group from an a-amino
acid to an a-keto acid, catalyzed by a transam-
inase. In biosynthetic reactions (a-amino
acicbi is often glutamate, with its carbon
skeleton producing a-ketoglutarate [= (a-keto
acid)iT (a-keto acid)2 represents the precur-
sor of a newly formed acid, (a-amino acid)2-

(a)

NH 4 ©-^

(b)

u-Ketoglutarate Amino acid

Glutamine u-Ketoglutarate Amino acid

Glutamate

dehydrogenase

Glutamate

Transaminase

u-Keto acid

Glutamine

synthetase

Glutamate

synthase

Glutamate

Transaminase

u-Keto acid

▲ Figure 17.8

Assimilation of ammonia into amino acids, (a) The glutamate dehydrogenase pathway, (b) Combined action of glutamine synthetase
and glutamate synthase under conditions of low NH 4 © concentration.

520

CHAPTER 17 Amino Acid Metabolism

17.3 Synthesis of Amino Acids

We now turn our attention to the origins of the carbon skeletons of amino acids. Figure 17.9
shows how the biosynthesis pathways leading to the 20 common amino acids are related to
other metabolic pathways. Note that 1 1 of the 20 common amino acids are synthesized from
intermediates in the citric acid cycle. The others require simple precursors that we have en-
countered in previous chapters.

A. Aspartate and Asparagine

Oxaloacetate is the amino group acceptor in a transamination reaction that produces
aspartate (Figure 17.10). The enzyme that catalyzes this reaction is aspartate transami-
nase (L-aspartate:2-oxoglutarate aminotransferase). Asparagine is synthesized in most
species by an ATP- dependent transfer of the amide nitrogen of glutamine to aspartate
in a reaction catalyzed by asparagine synthetase. In some bacteria, asparagine synthetase
catalyzes the formation of asparagine from aspartate using ammonia instead of gluta-
mine as the source of the amide group. This reaction is similar to the reaction catalyzed
by glutamine synthetase.

Some asparagine synthetases can use either ammonia or glutamine as the substrate.
These enzymes use NH 4 © at the primary reaction site but they have a second site that
catalyzes hydrolysis of glutamine and release of NH 4 ®. The NH 4 ® intermediate diffuses
through a tunnel in the protein that connects the two active sites. This example of molec-
ular channeling ensures that the hydrolysis of glutamine is tightly coupled to the forma-
tion of asparagine and it prevents the accumulation of NH 4 ® in the cell. There are many
examples of molecular tunnels that facilitate the channeling of NH 4 ® (see Box 18.2).

B. Lysine, Methionine, and Threonine

Aspartate is the precursor for synthesis of lysine, methionine, and threonine
(Figure 17.1 1). The first step in the pathway is the phosphorylation of aspartate in a re-
action catalyzed by aspartate kinase. In the second step, aspartyl phosphate is converted
to aspartate /3-semialdehyde. This second reaction is catalyzed by aspartate semialde-
hyde dehydrogenase. These two enzymes are present in bacteria, protists, fungi, and
plants but they are missing in animals. Consequently, animals are not able to synthesize
lysine, methionine, and threonine (see Box 17.3).

The first two reactions leading to aspartate /3- semialdehyde are common to the for-
▼ Figure 17.9 mation of all three amino acids. In the branch leading to lysine, pyruvate is the source of

Biosynthesis of amino acids, showing the carbon atoms added to the skeleton of aspartate /3- semialdehyde and glutamate is the

connections to glycolysis/gluconeogenesis
and the citric acid cycle.

Cysteine

Glycine

Cysteine

Isoleucine

Asparagine

Lysine

Methionine

^^.Threonine

Glucose 6-phosphate > Ribose 5-phosphate > Histidine

Serine < 3-Phosphoglycerate

Erythrose 4-phosphate

Pyruvate -

Phosphoenolpyruvate

Alanine
>| Valine
Leucine

u-Ketoglutarate — > Glutamate

Phenylalanine

Tryptophan

Arginine

Glutamine

Proline

Tyrosine

17.3 Synthesis of Amino Acids 521

coo 0

u-Ketoglutarate

coo^

^ 1

c=o

Glutamate/

HoN — CH
1

cn 2

Aspartate transaminase

cn 2

# \ o

0 O 0

o y x o c

Oxaloacetate

Aspartate

PPi

ATP AMP

'Asparagine^
synthetase

Glutamine

Glutamate

coo 0

H 3 N — CH

Asparagine

◄ Figure 17.10
Synthesis of aspartate and
asparagine.

source of the s-amino group. Lysine is produced by an entirely different route in yeast
and some algae.

Homoserine is formed from aspartate /3-semialdehyde. It is a branch point for the
formation of threonine and methionine. Threonine is derived from homoserine in two
steps, one of which requires PLP. In the methionine pathway homoserine is converted to
homocysteine in three steps. The sulfur atom of homocysteine then accepts a methyl
group derived from 5-methyltetrahydrofolate forming methionine. The enzyme that cat-
alyzes this reaction is homocysteine methyltransferase, one of the few enzymes that re-
quires cobalamin (Section 7.12). Homocysteine methyltransferase is found in mammals
but its activity is low and the supply of homocysteine is limited. Therefore, methionine
remains an essential amino acid in mammals due primarily to the absence of the first
two enzymes in the pathway.

C. Alanine, Valine, Leucine, and Isoleucine

Pyruvate is the amino group acceptor in the synthesis of alanine by a transamination re-
action (Figure 17.12). Pyruvate is also a precursor in the synthesis of the branched chain
amino acids valine, leucine, and isoleucine. The first step in the branched chain pathway
is the synthesis of a-ketobutyrate from threonine.

Pyruvate combines with a-ketobutyrate in a series of three reactions leading to the
branched chain intermediate a-keto-/3-methylvalerate. This intermediate is converted
to isoleucine in a transamination reaction. Note the similarity between the structures of
u-ketobutyrate and pyruvate. The same enzymes that catalyze the synthesis of u-keto-/3-
methylvalerate also catalyze the synthesis of a-ketoisovalerate by combining two molecules
of pyruvate instead of one molecule of pyruvate and one molecule of a-ketobutyrate.
a-Ketoisovalerate is converted directly to valine by valine transaminase — the same en-
zyme catalyzes the synthesis of isoleucine from a-keto-/3-methylvalerate (Figure 17.13).
These pathways illustrate an important point, namely that some enzymes recognize sev-
eral different but similar substrates. At some point in the future the eukaryotic genes for
these enzymes might be duplicated and each of the two copies would evolve to become
specific for either the isoleucine or valine pathways. If this happened, it would be an exam-
ple of pathway evolution by gene duplication and divergence (Section 10. 2D). We see

▲ Asparaginases, (a) Escherichia coli [PDB
INNS] (b) Erwinia chrysanthemi [PDB 107J]

BOX 17.1 CHILDHOOD ACUTE LYMPHOBLASTIC LEUKEMIA CAN BE TREATED WITH ASPARAGINASE

Acute lymphoblastic leukemia (ALL) is caused by the prolif-
eration of malignant T-cell lymphoblasts due, in most cases,
to a mutation caused by mistakes in genetic recombination
during the activation of T-cell receptor genes. Malignant
lymphoblasts have reduced levels of asparagine synthetase
and are unable to synthesize enough asparagine to support
their rapid growth and proliferation. Unlike normal cells,
they must obtain asparagine from the blood plasma.

This cancer can be successfully treated with injections of
asparaginase, an E. coli enzyme that breaks down asparagine
in the plasma (Section 17. 6 A). The malignant cells die in the
absence of an available source of asparagine. Treatment with

asparaginase alone causes remission in 50% of all cases of
childhood acute lymphoblastic leukemia and the success rate
is even higher when the enzyme treatment is combined with
other chemotherapy. The primary cause of resistance to the
treatment is due to increased expression of asparagine syn-
thetase in the cancer cells.

Patients often develop antibodies to the E. coli enzyme
during treatment. Switching to the homologous enzyme
from Erwinia chrysanthemi is often effective because the
amino acid side chains on the surface of the two proteins are
different. Antibodies directed against one enzyme usually
don’t recognize the other.

522 CHAPTER 17 Amino Acid Metabolism

COO'

,©

coo

HoN — CH

CH,

ATP ADP

H 3 N — CH

CH,

NADPH NADP©
+ +

H 0 P;

PUN -

COO G

Aspartate

kinase

\>P0 3 ®
f3 - Aspartyl phosphate

Aspartate

semialdehyde

dehydrogenase

COO'

-CH

,©

coo'

,©

8 reactions

O H
Aspartate /3-semialdehyde

NAD(P)H, H©
NAD(P)©^

H,N — CH

(CH 2 ) 4

©nh 3

Lysine

COO

,©

COO'

I©

coo'

,©

coo'

HoN — CH

cn,

ch 3

Methionine

«-

HoN — CH

CH.

CH 2

Homocysteine

3 reactions

HoN — CH

CHo

ATP ADP H 2 0 Pj

V ± » ^ ,

(PLP)

ch 2 oh

Homoserine

HoN— CH

h 3 c oh

Threonine

▲ Figure 17.1 1

Biosynthesis of lysine, threonine, and methionine
from aspartate.

coo©

C = 0
1

coo©

COO©

ch 2

C = 0
1

ch 3

a:-Ketobutyrate

Pyruvate

3 reactions \

/ N

/ 3 reactions

Amino a-Keto
acid acid

Transamination

(PLP)

COO'

,©

HoN — CH

CH 3

Alanine

H,C

s co 2

Sco 2

coo©

C = 0
1

-U-

ch 2

h 3 c

ch 3

4 reactions

CHo

a-Ketoisovalerate

o;-Keto-/3-methylvalerate

coo©

coo c

H 3 N — CH

H 3 N — CH
1

H 3 N — CH

ch 2

/ \

1 2

h 3 c ch 2

h 3 c ch 3

Valine

/ \

ch 3

H 3 C CH

Isoleucine

Leucine

◄ Figure 17.12

Biosynthesis of alanine, isoleucine,
valine, and leucine.

17.3 Synthesis of Amino Acids 523

many examples of pathway evolution by gene duplication involving enzymes of amino
acid metabolism (see below). The basic requirement is that in the early stages the same
enzyme can catalyze two similar reactions and that is what we see in the isoleucine and
valine synthesis pathways.

The carbon skeleton of a-ketoisovalerate is lengthened by one methylene group to
form leucine in a pathway that branches from the valine biosynthetic pathway. Two of
the enzymes in this pathway are homologous to aconitase and isocitrate dehydrogenase
in the citric acid cycle lending support to the idea that citric acid cycle enzymes evolved
from preexisting enzymes required for amino acid biosynthesis (Section 13.8).

D. Glutamate, Glutamine, Arginine, and Proline

We have seen how glutamate and glutamine are formed from the citric acid cycle inter-
mediate a-ketoglutarate (Section 17.2B). The carbon atoms of proline and arginine also
come from a-ketoglutarate, via glutamate. Proline is synthesized from glutamate by a
four-step pathway in which the 5-carboxylate group of glutamate is reduced to an alde-
hyde. The glutamate 5 -semialdehyde intermediate undergoes nonenzymatic cyclization
to a Schiff base, 5-carboxylate, that is reduced by a pyridine nucleotide coenzyme to
produce proline (Figure 17.14).

The pathway to arginine is similar in most species except that the a-amino group
of glutamate is acetylated before the aldehyde is formed. This step prevents the cycliza-
tion that occurs in the synthesis of proline. The N-acetylglutamate 5-semialdehyde
intermediate is then converted to N-acetyl ornithine and ornithine. In mammals, gluta-
mate 5 -semialdehyde is transaminated to ornithine and ornithine is converted to argi-
nine by the reactions of the urea cycle (Section 17.7).

u-Ketobutyrate

Pyruvate

Acetohydroxy acid

synthase

Acetohydroxv acid isomeroreductase
Dihydroxy acid dehydratase
Valm^ aminotransferase

Isoleucine Valine

▲ Figure 17.13

The isoleucine and valine synthesis pathways
share four enzymes.

E. Serine, Glycine, and Cysteine

Three amino acids — serine, glycine, and cysteine — are derived from the glycolytic/
gluconeogenic intermediate 3-phosphoglycerate. Serine is synthesized from 3-phospho-
glycerate in three steps (Figure 17.15). First, the secondary hydroxyl substituent of

Figure 17.14 ►

Conversion of glutamate to proline and arginine.

coo°

H,N — CH
1

h 3 c— c —

NH — CH

F -

— > >

i H;

°\

H 0

Glutamate

A/-Acety I g I uta m ate
5-semialdehyde

COO

0 / c ^

h 2 n ch 2

\ /

h 2 c — ch 2

Proline

COO 0

m CH
HN CH 2

w /

HC — CH 2

A^Pyrroline

5-carboxylate

coo°

H 3 N— CH

H 3 N — CH

CH,

> >

CH,

ch 2

/ \

H 0

@ nh 3

Glutamate

Ornithine

5-semialdehyde

/ %©

H 2 N NH

Arginine

524

CHAPTER 17 Amino Acid Metabolism

coo°

H — C — OH

NAD® NADH + H®

,©

ch 2 opo 3

3-Phosphoglycerate

dehydrogenase

COO'

c = o

,© Glutamate u-Ketoglutarate

Phosphoserine
transaminase
2 uru 3 (PLP)

3-Phosphohydroxypyruvate

COO'

H 3 N — CH

,©

ch,opo 3 ©

ch 2 opo 3

3-Phosphoserine

H 2 0 P;

L 1 »

3-Phosphoserine
@ phosphatase

COO 1

HoN — CH

CH 2 OH

Serine

▲ Figure 17.15
Biosynthesis of serine.

3-phosphoglycerate is oxidized to a keto group, forming 3-phosphohydroxypyruvate.
This compound undergoes transamination with glutamate to form 3-phosphoserine
and a-ketoglutarate. Finally, 3-phosphoserine is hydrolyzed to give serine and Pj.

Serine is a major source of glycine via a reversible reaction catalyzed by serine hy-
droxymethyltransferase (Figure 17.16). In plant mitochondria and bacteria, the flux
through this reaction is toward serine providing a route to serine that differs from that
in Figure 17.15. The serine hydroxymethyltransferase reaction requires two cofactors:
the prosthetic group PLP and the cosubstrate tetrahydrofolate.

The biosynthesis of cysteine from serine occurs in two steps (Figure 17.17). First,
an acetyl group from acetyl CoA is transferred to the /3-hydroxyl substituent of serine,
forming O-acetylserine. Next, sulfide (S©) displaces the acetate group, and cysteine is
formed.

Animals do not have the normal cysteine biosynthesis pathway shown in Figure 17.17.
Nevertheless, cysteine can still be synthesized in animals as a by-product of methionine
degradation (Section 17.6F). Serine condenses with homocysteine, an intermediate in
the degradation of methionine. The product of the condensation reaction, crystathionine,
is cleaved to a-ketobutyrate and cysteine (Figure 17.18).

v Figure 17.16
Biosynthesis of glycine.

coo®

H 3 N — CH

I +

ch 2 oh

Serine

F. Phenylalanine, Tyrosine, and Tryptophan

The key to elucidation of the pathway for aromatic amino acid synthesis was the obser-
vation that some bacteria with single-gene mutations require as many as five com-
pounds for growth: phenylalanine, tyrosine, tryptophan, p-hydroxybenzoate, and
p-aminobenzoate. These compounds all contain an aromatic ring. The inability of
these mutants to grow without these compounds is reversed when shikimate is pro-
vided indicating that shikimate is an intermediate in the biosynthesis of all these
aromatic compounds.

Chorismate, a derivative of shikimate, is a key branch-point intermediate in aro-
matic amino acid synthesis. The pathway to shikimate and chorismate (Figure 17.19)
begins with condensation of phosphoenolpyruvate and erythrose 4-phosphate to form
a seven- carbon sugar derivative and Three additional steps, including cyclization, are

required to produce shikimate. The pathway from shikimate to chorismate involves
phosphorylation of shikimate, addition of a three-carbon group from phospho-
enolpyruvate, and dephosphorylation. Pathways from chorismate lead to phenylala-
nine, tyrosine, and tryptophan. Animals do not have the enzymes of the chorismate
pathway. They cannot synthesize chorismate and, consequently, cannot synthesize any
of the aromatic amino acids.

A branched pathway leads from chorismate to phenylalanine or tyrosine (Figure 17.20).
In phenylalanine synthesis in E. coli , a bifunctional chorismate mutase-prephenate dehy-
dratase catalyzes the rearrangement of chorismate to produce prephenate, a highly reac-
tive compound. Next, the enzyme catalyzes the elimination of a hydroxide ion and C0 2

Tetrahydrofolate

H 2 0

Serine

hydroxymethyltransferase

(PLP)

coo®

h 3 n — ch 2

Glycine

H 2 C — N— R

1 10

5,10-Methylenetetrahydrofolate

17.3 Synthesis of Amino Acids 525

coo°

◄ Figure 17.17

0 1

Acetyl CoA HS-CoA

+ H® Acetate

0 1

Biosynthesis of cysteine from serine

H 3 N — CH

L 7 .

H,N — CH

L 7 .

H 3 N — CH

in many bacteria and plants.

ch 2

Serine

acetyltransferase

H,C O

1 II

O-Acetylserine

sulfhydrylase

ch 2

(PLP)

o— c — ch 3

Serine

O-Acetylserine

Cysteine

from prephenate to form the fully aromatic product phenylpyruvate that is then
transaminated to phenylalanine.

A similar bifunctional chorismate mutase-prephenate dehydrogenase catalyzes the
formation of prephenate and then 4-hydroxyphenylpyruvate in the tyrosine branch.
The intermediate undergoes transamination to form tyrosine. Several bacteria and
some plants follow the same pathways from chorismate to phenylalanine and tyrosine
as E. coli although their chorismate mutase and prephenate dehydratase or prephenate
dehydrogenase activities are on separate polypeptide chains. Some other bacteria use al-
ternate pathways in which prephenate is first transaminated and then decarboxylated.

The biosynthesis of tryptophan from chorismate requires five enzymes. In the first
step, the amide nitrogen of glutamine is transferred to chorismate. Subsequent elimina-
tion of the hydroxyl group and the adjacent pyruvate moiety of chorismate produces
the aromatic compound anthranilate (Figure 17.21). Anthranilate accepts a phosphori-
bosyl moiety from PRPP. Rearrangement of the ribose, decarboxylation, and ring clo-
sure generate indole glycerol phosphate.

The final two reactions of tryptophan biosynthesis are catalyzed by tryptophan
synthase (Figure 17.22). In some organisms, the two independent catalytic domains of
tryptophan synthase are contained on a single polypeptide chain but in some species
the enzyme contains two types of subunits in an a 2 (3 2 tetramer. The a subunit, or do-
main, catalyzes the cleavage of indole glycerol phosphate to glyceraldehyde 3 -phosphate
and indole. The (3 subunit, or domain, catalyzes the condensation of indole and serine
in a reaction that requires PLP as a cofactor. The indole produced in the reaction cat-
alyzed by the a subunit of ol 2 (3 2 tetramers is channeled (i.e., transferred directly) to the
active site of the (3 subunit. When the three-dimensional structure of tryptophan syn-
thase from Salmonella typhimurium (an organism whose tryptophan synthase has the
OL 2 f3 2 oligomeric structure) was determined by X-ray crystallography, a tunnel joining
the a and (3 active sites was discovered. The diameter of the tunnel matches the molecu-
lar dimensions of indole, so passage of indole through the tunnel would explain why

COO

,©

coo (

,©

h 3 n — ch

r 2

Serine

CH 2

^ H2 ©

HC — NH,

Cystathionine

/3-synthase

(PLP)

H,0

COO°

H 3 N — CH

CH 2

T 2 ©

HC— NH,

COO 0

Homocysteine

Cystathionine

y-lyase

(PLP)

H?0 NH

coo°

Cystathionine

H 3 N — CH

ch 2

Cysteine

cn 3

ch 2

c = o

1 0

coo°

u-Ketobutyrate

◄ Figure 17.18

Biosynthesis of cysteine in mammals.

526 CHAPTER 17 Amino Acid Metabolism

coo 0

c— OPO3®

ch 2

Phosphoenolpyruvate

coo°

4 reactions

Shikimate

kinase

coo 0

H — C — OH

CH 2 0P0 3 ©
Erythrose 4-phosphate

Shikimate

5-phosphate

PEP

5-Enolpyruvylshikimate-
3-phosphate synthase
(EPSP synthase)

coo 0

Chorismate

synthase

coo 0

5-Enolpyruvylshikimate

3-phosphate

a Figure 17.19

Synthesis of shikimate and chorismate.

Figure 17.20 ►

Biosynthesis of tryptophan, phenylalanine, and
tyrosine from chorismate in E. coli.

coo 0

Chorismate

®NhU

Chorismate mutase

Phenylpyruvate

co 2 ,oh°

♦ — I

Prephenate

dehydratase

(PLP)

Glutamate

a-Ketoglutarate

Prephenate

NADH

NAD®

« >2

J—

Prephenate

dehydrogenase

4-Hydroxyphenylpyruvate

Glutamate

a-Ketoglutarate

(PLP)

Phenylalanine

©NH 3

— CH — COO®

Tyrosine

17.3 Synthesis of Amino Acids

527

indole does not diffuse away. This was one of the earliest examples of metabolic chan-
neling (Section 5.10). Up until quite recently there were only a few other examples and
the phenomenon was thought to be rare. The huge increase in structural and genomic
studies has revealed many more examples — including half a dozen in this chapter alone.

G. Histidine

The ten-step pathway for the biosynthesis of histidine in bacteria begins with a condensa-
tion between the six-membered ring of ATP and a ribose derivative, phosphoribosyl
pyrophosphate (PRPP) (Figure 17.23). The six-membered ring of the adenine moiety is
then cleaved and glutamine donates a nitrogen atom that is incorporated via cyclization

▲ Figure 17.21
Anthranilate.

OH OH

1 1 ©
CH — CH — CH 2 0P0 3 ^

Indole glycerol phosphate

Glyceraldehyde

3-phosphate

■>

Tryptophan

synthase

(a)

®nh 3

1 ©

HOCH 2 — CH — COO°

Indole

Serine

H 2 0

L 7,

Tryptophan

synthase

03 )

(PLP)

▲ Figure 17.22

Reactions catalyzed by tryptophan synthase.

v Figure 17.23

Synthesis of histidine from phosphoribosyl pyrophosphate (PRPP) and ATP.

Histidine is derived from PRPP (5 C atoms), the purine ring of ATP
(1 N and 1 C), glutamine (1 N), and glutamate (1 N).

a-D-Phosphoribosyl pyrophosphate
(PRPP)

N ^ N -- R i bose-®-®-®

3 reactions

| Hz

HC — NH 3

COO'

■©

Glutamine

Glutamate

-N

5 reactions

^Ribose-®
NH,

H 2 N

Aminoimidazole carboxamide
ribonucleotide

-> Purine biosynthesis

H — C — OH

CH 2 OPO:

Histidine

Imidazole glycerol
phosphate

528 CHAPTER 17 Amino Acid Metabolism

BOX 17.2 GENETICALLY MODIFIED FOOD

The chorismate pathway is an effective target for herbicides
since compounds that specifically block this pathway in plants
will have no effect on animals. One of the most effective gen-
eral herbicides is glyphosate. Glyphosate inhibits the enzyme
5-enolpyruvylshikimate-3-phosphate synthase (EPSP syn-
thase) by acting as a competitive inhibitor of PEP binding
(Section 5.7A).

Glyphosate is the active ingredient in Roundup®, a her-
bicide that kills all plants. It is used to remove weeds from
driveways and stone pathways. Although it is cheap and ef-
fective as a weed killer, glyphosate cannot be used to spray ac-
tively growing food crops since it indiscriminately kills all
plants, including the crop!

®o 3 p— ch 2 — nh— ch 2 — coo 0

Glyphosate

(N-(phosphonomethyl) glycine)

Resistant versions of EPSP synthase have been identified
in many species of bacteria. The enzyme from strain CP4 of
Agrobacterium sp. has been genetically modified to remain
fully active in the presence of high concentrations of glyphosate.
The gene for this bacterial CP4-EPSP synthase was patented
and then introduced into soybeans creating a genetically
modified plant that is resistant to glyphosate. The new strain

of soybeans is marketed by Monsanto as Roundup Ready®
soybeans. Farmers who grow crops of Roundup Ready®
soybeans are able to spray them with Roundup® (also sold by
Monsanto) to kill weeds. The economic advantages to farm-
ers are significant. Most of the soybeans currently grown in
North America are genetically modified.

Other Roundup Ready® crop plants are now available.
Versions of corn, cotton, and canola are widely used.

▲ E. coli 5-enolpyruvylshikimate-3-phosphate synthase with a molecule
of glyphosate bound to the active site. [PDB 2AAY]

KEY CONCEPT

Metabolic channeling evolves to improve
kinetic efficiency.

into the imidazole ring of the product, imidazole glycerol phosphate. Most of the carbon
and nitrogen atoms of ATP are released as aminoimidazole carboxamide ribonucleotide,
an intermediate in purine biosynthesis (Section 18.1). This metabolite then can be recycled
into ATP. Imidazole glycerol phosphate undergoes dehydration, transamination by gluta-
mate, hydrolytic removal of its phosphate, and oxidation from the level of a primary al-
cohol to that of a carboxylic acid in two sequential NAD® -dependent steps forming
histidine.

▲ Tryptophan synthase from Salmonella typhimurium. The substrate indole glycerol phosphate is
shown as a space-filling molecule bound to the a subunits. The cofactor PLP is bound to the
/3 subunits. The enzyme contains a channel leading from the indole glycerol phosphate binding
site to the PLP reaction site. [PDB 1Q0Q]

17.4 Amino Acids as Metabolic Precursors 529

BOX 17.3 ESSENTIAL AND NONESSENTIAL AMINO ACIDS IN ANIMALS

Humans and other animals do not possess the enzymes re-
quired for the synthesis of all amino acids. Those that cannot
be synthesized are, therefore, essential components of the
human diet. As a general rule, the pathways that have been
lost are the ones with the most steps. A crude measure of the
complexity of a pathway is the number of moles of ATP (or
its equivalent) required in a pathway.

The table shows the correlation between the expense of a
particular pathway and whether an amino acid is essential. The
amino acids are grouped according to their common precur-
sors as described in the previous sections. Note that lysine, me-
thionine, and threonine are derived from a common precursor
(Section 17.3B). All three amino acids are essential because ani-
mals cannot synthesize the precursor. Valine, leucine, and
isoleucine are essential because animals lack the key enzymes
that all three biosynthesis pathways share (Section 17.3C).

a Moles of ATP required includes ATP used for synthesis of precursors and conversion
of precursors to products.
b Essential in some mammals.

c Cysteine can be synthesized from homocysteine and homocysteine is a degradation
product of methionine. The biosynthesis of cysteine depends on an adequate supply
of methionine in the diet.

d Tyrosine can be synthesized from the essential amino acid phenylalanine.

Energy requirements for biosynthesis of amino acids

Moles of ATP required per
Amino acids mole of amino acid produced 21

Nonessential

Essential

Aspartate

Asparagine

22-24

Lysine

50 or 51

Methionine

Threonine

Alanine

Valine

Leucine

Isoleucine

Glutamate

Glutamine

Arginine

44 b

Proline

Serine

Glycine

Cysteine

1 9 C

Phenylalanine

Tyrosine

Tryptophan

62 d

Histidine

17.4 Amino Acids as Metabolic Precursors

The primary role of amino acids is to serve as substrates for protein synthesis. In this
role, newly synthesized amino acids are activated by covalent attachment to tRNA and
the pool of aminoacyl-tRNAs is used as the substrate for polypeptide synthesis by the
protein synthesis machinery. We devote an entire chapter to this fundamentally impor-
tant biosynthesis pathway (Chapter 22).

Some amino acids are essential precursors in other biosynthesis pathways. The list is
long and if s impossible to mention every pathway. Some important regulatory amines were
described in Section 3.3 (histamine, GABA, epinephrine, thyroxine). The important role of
methionine in the synthesis of S-adenosylmethionine will be described in Section 17.6F.

A. Products Derived from Glutamate, Glutamine, and Aspartate

Weve already seen that glutamate and glutamine are important players in nitrogen assim-
ilation. In addition, glutamate and aspartate are amino group donors in many transami-
nation reactions. We will see that glutamate and aspartate are required in the urea cycle.
Glutamine and aspartate are also required as precursors in both purine biosynthesis (Sec-
tion 18.1) and pyrimidine biosynthesis (Section 18.3). Recall that synthesis of biologically
active tertahydrofolate involves addition of up to six glutamate residues to the tetrahydro-
folate moiety (Section 7.10).

Phenylalanine

tRNAPhe

B. Products Derived from Serine and Glycine

Serine and glycine are metabolic precursors of many other compounds (Figure 17.24).
The role of serine in lipid biosynthesis has already been described in the previous chapter.

▲ Phenylanyl-tRNA Phe . Most newly synthe-
sized amino acids are rapidly attached to
their corresponding tRNAs and used in
protein synthesis. [PDB 1TTT]

530 CHAPTER 17 Amino Acid Metabolism

Figure 17.24 ►

Compounds formed from serine and glycine.

▲ Nitric oxide

In 1998 Robert F. Furchgott, Louis J.
Ignarro, and Ferid Murad were awarded
the Nobel Prize in Physiology or
Medicine “for their discoveries
concerning nitric oxide as a signaling
molecule in the cardiovascular system.”

Phosphatidylcholines

Sphingol ipids

Phosphatidylethanolamines

Sphinganine

■ Phosphatidylserines

Tetrahydrofolate

Deoxythymidylate < Methylene- 4 —

tetrahydrofolate

Serine

Cysteine

Methyl- Purines

tetrahydrofolate

Glycine

Glutathione

Methionine

Bile salts \ Glyoxylate

Succinyl

CoA

Porphobilinogen Creatine phosphate

Porphyrins

Chlorophyll Heme Cobalamin

Glycine and succinyl CoA are the main precursors in the porphyrin pathway leading to
heme and chlorophyll. Glycine is also required in purine biosynthesis (Section 18.1).

The conversion of serine to glycine is coupled to the synthesis of methylene
tetrahydrofolate. Tetrahydrofolate derivatives are important in many reactions that cat-
alyze transfer of one-carbon units (Section 17.10). One of the most important of these
reactions is the synthesis of deoxythymidylate (Figure 18.15).

C. Synthesis of Nitric Oxide from Arginine

One of the more interesting examples of amino acids as metabolic precursors is the role
of arginine as substrate for synthesis of nitric oxide, an unstable gaseous derivative of
nitrogen with an odd number of electrons (*N = O). Although it is a reactive free radical
and potentially toxic, nitric oxide is physiologically important — so important, in fact,
that it was named the 1992 “Molecule of the Year” by the journal Science. As a gas, NO
can diffuse rapidly into cells. It exists in vivo for only a few seconds because nitric oxide
in aqueous solution reacts rapidly with oxygen and water to form nitrates and nitrites.

An enzyme found in mammals, nitric oxide synthase, catalyzes the formation of ni-
tric oxide and citrulline from arginine (Figure 17.25). The reaction requires the cofac-
tors NADPH, FMN, FAD, a cytochrome P450, and tetrahydrobiopterin (Section 7.10).
The mechanism of action of tetrahydrobiopterin in this reaction has not yet been eluci-
dated but it appears to be a reducing agent needed for the hydroxylation of arginine.
Nitric oxide synthase is present in two forms, a constitutive (i.e., constantly synthesized)
calcium-dependent form in brain and endothelial cells and an inducible (i.e., variably
synthesized) calcium-independent form in macrophages (a type of white blood cell).

Nitric oxide is a messenger molecule that binds to a soluble guanylyl cyclase and
stimulates the formation of cyclic GMP (Section 9.12B). It has several functions; for
example, when macrophages are stimulated, they synthesize nitric oxide. The short-lived
nitric oxide free radical is one of the weapons used by macrophages to kill bacteria and
tumor cells. Nitric oxide may interact with superoxide anions (*0 2 ® ) to form more toxic
reactants that account for the cell-killing activity.

17.5 Protein Turnover 531

©nh 2

2 0 2 , 3 e 0 2 OH 0

nh 2

◄ Figure 17.25

Conversion of arginine to nitric oxide and

c — NH,

citrulline. NADPH is the source of the three

V V .

electrons.

Nitric oxide synthase

i Hi

ch 2

{ H >

CH— COO 0

CH — COO 0

©nh 3 ©nh 3

Arginine Citrulline

Nitric oxide synthase is also present in the cells that line blood vessels. Under cer-
tain conditions, nitric oxide is produced and diffuses to the smooth muscle cells of the
vessels, causing them to relax and lower blood pressure. Hypertension and heart failure
involve impaired relaxation of blood vessels. Nitroglycerin, used to dilate coronary ar-
teries in the treatment of angina pectoris, exerts its effect by virtue of its metabolic con-
version to nitric oxide.

Nitric oxide also functions as a neurotransmitter in brain tissue. Abnormally high
amounts of nitric oxide formed during a stroke appear to kill some neurons in the same
way macrophages kill bacteria. Administering an inhibitor of nitric oxide synthase to an
animal produces some protection from stroke damage. One role of nitric oxide as a neu-
rotransmitter is to stimulate erection of the penis. Sildenafil, the active ingredient in Viagra,
is a drug used to alleviate erectile dysfunction. Sildenafil is a phosphodiesterase inhibitor
that blocks the hydrolysis of cyclic GMP and therefore prolongs the stimulatory effect of
nitric oxide. Tadalafil (Cialis) and vardenafil (Levitra) inhibit the same enzyme.

D. Synthesis of Lignin from Phenylalanine

Lignin (Figure 17.26) is a series of complex polymers synthesized from phenylalanine. It
is a major component of wood in flowering plants and may be the second most abun-
dant biopolymer on the planet (after cellulose). Lignin cannot be broken down during
digestion so in spite of the fact that animals ingest huge amounts of lignin it is metabol-
ically inert. The only species that can break it down are various fungi that degrade fallen
trees in the forest.

E. Melanin Is Made from Tyrosine

Melanin is a dark pigment found in bacteria, fungi, and animals. In humans it is re-
sponsible for skin color and hair color. Melanin is also the main component of the ink
released by a frightened octopus.

The structure of melanin (eumelanin) is complex but the precursors are well
known and the enzymes required in the pathway have been identified in a number of
species. The first steps involve the conversion of L-tyrosine to l-DOPA and
L-dopaquinone (Figure 17.27).

17.5 Protein Turnover

One might assume that only growing or reproducing cells would require new protein mol-
ecules (and therefore a supply of amino acids) but this is not the case. Proteins are continu-
ally synthesized and degraded in all cells, a process called turnover. Individual proteins turn
over at different rates. Their half-lives can vary from a few minutes to several weeks but the
half-life of a given protein in different organs and species is generally similar. Rapid protein
turnover ensures that some regulatory proteins are degraded so that the cell can respond
to constantly changing conditions. Such proteins have evolved to be relatively unstable.

CH 3

▲ Sildenafil. Sildenafil is the active ingredient
in Viagra®.

▲ Octopus ink is mostly melanin.

532 CHAPTER 17 Amino Acid Metabolism

Figure 17.26 ►

Lignin. This is one of many possible struc-
tures of plant lignin.

h 2 coh

HCOR

H 3 CO

och 3

▲ Rotting wood. This mushroom is grow-
ing on rotting wood in a deciduous forest.
Fungi are the only organisms that pro-
duce enzymes for breaking down lignin.

The rate of hydrolysis of a protein can be inversely related to the stability of its tertiary
structure. Misfolded and unfolded proteins are quickly degraded (Section 4.10).

Some proteins are degraded to amino acids through lysosomal hydrolysis (in eu-
karyotic cells). Vesicles containing material to be destroyed fuse with lysosomes, and
various lysosomal proteases hydrolyze the engulfed proteins. The lysosomal enzymes
have broad substrate specificities so all the trapped proteins are extensively degraded.

Some proteins have very short half-lives because they are specifically targeted for
degradation. Abnormal (mutated) proteins are also selectively hydrolyzed. The pathway
for the selective hydrolysis of these proteins in eukaryotic cells requires the protein
ubiquitin. Side-chain amino groups of lysine residues in the target protein are co-
valently linked to the C-terminus of ubiquitin in a complex pathway that involves

17.5 Protein Turnover 533

COO G

coo 0

l 3 N — C — H

H 3 N — C — H

b -

-u —
1

V^OH

Tyrosine

L-DOPA

◄ Figure 17.27

Synthesis of eumelanin from tyrosine and
l-DOPA.

coo 0

H 3 N— C — H

L-Dopaquinone

Eumelanin

ubiquitin- activating enzyme (El), ubiquitin- conjugating enzyme (E2), and ubiquitin-
protein ligase (E3). This pathway is coupled to ATP hydrolysis — one ATP molecule is
hydroylzed for every ubiquitin molecule attached to the target protein. The ubiquiti-
nated protein is hydrolyzed to peptides by the action of a large multiprotein complex
called the proteasome (or proteosome) (Figure 17.28). This process occurs in both the
cytosol and the nucleus. Other proteases catalyze hydrolysis of the resulting peptides.
ATP is required to assemble the proteasome and to hydrolyze the ubiquitinated protein.
Before this pathway was discovered there was no explanation for the surprising observa-
tion that the degradation of many proteins requires ATP. (Recall from Section 2.6 that
hydrolysis of a peptide bond is a thermodynamically favorable reaction.)

▲ Ubiquitin [Homo sapiens). Ubiquitin is a
small, highly conserved, eukaryotic protein
used as a marker that targets proteins for
degradation. [PDB 1UBI]

Aaron Ciechanover (1947-), Avram
Hershko (1937-), and Irwin Rose
(1926-) won the 2004 Nobel Prize in
Chemistry “for the discovery of ubiquitin-
mediated protein degradation.”

Ubiquitinated protein

▲ Figure 17.28

Ubiquitination and hydrolysis of a protein. Ubiquitination enzymes catalyze the attachment of numerous molecules of ubiquitin to the protein targeted
for degradation. The proteasome catalyzes ATP-dependent hydrolysis of the substituted protein, releasing peptides and ubiquitin.

534 CHAPTER 17 Amino Acid Metabolism

BOX 17.4 APOPTOSIS— PROGRAMMED CELL DEATH

Apoptosis (often pronounced with the second p silent) is a
series of morphological changes in a cell that leads to its
death. The changes include a decrease in cell volume, damage
to the plasma membrane, swelling of mitochondria, and
fragmentation of chromatin. Surplus and harmful cells are
removed principally by the action of proteases.

Some cells die normally during development or in the
regulation of antibody production. Others die as a result of
diseases or from faulty apoptosis (as in some neurodegenera-
tive diseases). As a result of apoptosis, vesicles containing cel-
lular contents form and are engulfed by neighboring cells.
Some of the protein contents of the vesicles can be saved and
reused by the other cells.

All eukaryotes have a similar set of endogenous enzymes
responsible for cell death. These enzymes (first described as
being involved in apoptosis in 1993) include about a dozen

proteases called caspases — meaning cysteine- containing hy-
drolases that act on the carboxyl side of aspartate residues.

▲ Apoptosis. The drawing depicts vesicles from a dead apoptotic cell
(purple) being taken up by a white blood cell (green). [Courtesy of
the United States National Library of Medicine.]

17.6 Amino Acid Catabolism

Amino acids obtained from the degradation of endogenous proteins or from the diet can
be used for the biosynthesis of new proteins. Amino acids not needed for the synthesis of
proteins are catabolized in order to make use of their nitrogen and their carbon skele-
tons. The first step in amino acid degradation is often removal of the a -amino group.
Next, the carbon chains are altered in specific ways for entry into the central pathways of
carbon metabolism. We first consider the metabolic fates of the various carbon skeletons.
In the next section we examine the metabolism of the ammonia arising from amino acid
degradation. These catabolic pathways are present in all species but they are especially
important in animals since amino acids are a significant part of fuel metabolism.

Removal of the a-amino group of an amino acid occurs in several ways. The amino
acid usually undergoes transamination with a-ketoglutarate to form an cr-keto acid and
glutamate. The glutamate is oxidized to a-ketoglutarate and ammonia by the action of
mitochondrial glutamate dehydrogenase. The net effect of these two reactions is the re-
lease of a-amino groups as ammonia and the formation of NADH and cr-keto acids.
This is the reverse of the pathway shown in Figure 17.8A.

Amino acid + a-Ketoglutarate a-Keto acid + Glutamate

Glutamate + NAD© + H z O a-Ketoglutarate + NADH + H© + NH 4 ©

Sum: Amino acid + NAD© + H 2 0 a-Keto acid + NADH + H© + NH 4 © (17.4)

The amide groups of glutamine and asparagine are hydrolyzed by specific enzymes —
glutaminase and asparaginase, respectively — to produce ammonia and the correspon-
ding dicarboxylic amino acids glutamate and aspartate. Ammonia from amides and
amino groups that is not used in biosynthesis reactions is excreted.

Once the amino groups have been removed, the carbon chains of the 20 amino
acids can be degraded. Some are degraded to one of four citric acid cycle intermediates
while others are degraded to pyruvate, and still others to acetyl CoA or acetoacetate
(Figure 17.29). Each amino acid follows its own route to one or more of these seven
compounds.

While all these products can be oxidized to C0 2 and H 2 0 they can also have other
metabolic fates. Amino acids that are degraded to pyruvate or citric acid cycle intermediates
are called glucogenic because they can directly supply the pathway of gluconeogenesis.
Those that form acetyl CoA or acetoacetate can contribute to the formation of fatty
acids or ketone bodies and are called ketogenic. Some amino acids are both glucogenic

▲ Proteasome from yeast ( Saccharomyces
cerevisiae). (a) Side view. The complete pro-
teasome consists of two seven-member rings
of /3 subunits (blue) with their active pro-
tease sites on the interior of the cylinder.

The outer two rings have seven a subunits
(purple), (b) Top view. Ubiquinated proteins
enter the cylinder through a pore at the top
or bottom of the structure. [PDB 1FNT]

17.6 Amino Acid Catabolism

535

Tryptophan > Alanine

Cysteine

Asparagine > Aspartate

Glucose « Phosphoenolpyruvate <

Threonine
Serine

Glycine

Minor
pathway

^ 5,10-Methylenetetrahydrofolate

Phenylalanine

Tyrosine < —
Leucine
Lysine
Tryptophan

Acetoacetate

Pyruvate

Acetyl CoA

Oxaloacetate

Phenylalanine

-> Tyrosine
Aspartate

Fumarate

Citric

acid

cycle

u-Ketoglutarate

Fumarate

Succinyl CoA

Glucose

Isoleucine

Methionine

Valine

Threonine

Tyrosine < —

Isoleucine

Leucine

Lysine

Tryptophan

Threonine

Glutamate «

Phenylalanine

Arginine

Glutamine

Histidine

Proline

Key:

Glucogenic

Ketogenic

and ketogenic because different parts of their carbon chains form different products.
The distinction between glucogenic and ketogenic products is important in animals
since amino acids are significant fuel metabolites in the diet. Animals do not possess a
direct pathway leading from acetyl CoA to glucose and the production of excess acetyl
CoA stimulates formation of ketone bodies (Section 16.11). The distinction between
glucogenic and ketogenic products is less important in bacteria, protists, fungi, and
plants since they can convert acetyl CoA to oxaloacetate via the glyoxylate pathway
(Section 13.7). In these organisms, acetyl CoA is glucogenic.

In this section, we examine the pathways of amino acid degradation beginning
with the simplest routes. Our aim is to show how the carbon atoms of each amino acid
reach “glucogenic” metabolites (pyruvate and citric acid cycle intermediates) or “keto-
genic” metabolites (acetyl CoA and acetoacetate). The ultimate fates of these metabo-
lites depend on the species and are covered in earlier chapters.

▲ Figure 17.29

Degradation of amino acids. The carbon
skeletons of amino acids are converted to
pyruvate, acetoacetate, acetyl CoA, or citric
acid cycle intermediates.

A. Alanine, Asparagine, Aspartate, Glutamate, and Glutamine

Alanine, aspartate, and glutamate are synthesized by reversible transamination reactions
(Sections 17.3A,C,D). The breakdown of these three amino acids involves their re-entry
into the pathways from which their carbon skeletons arose. Alanine gives rise to pyruvate,
aspartate to oxaloacetate, and glutamate to a-ketoglutarate by reversal of the original
transamination reactions. All three amino acids are glucogenic since aspartate and gluta-
mate are converted to citric acid cycle intermediates and alanine is converted to pyruvate.

The degradation of both glutamine and asparagine begins with their hydrolysis to
glutamate and aspartate, respectively. Thus, glutamine and asparagine are both glucogenic.
The hydrolysis reactions are catalyzed by specific enzymes — asparaginase (Box 17.1)
and glutaminase.

B. Arginine, Histidine, and Proline

The pathways for the degradation of arginine, histidine, and proline converge on glutamate
(Figure 17.30). In the case of arginine and proline, the degradation pathways resemble
the biosynthesis pathways. Arginine degradation commences with the reaction catalyzed
by arginase. The ornithine produced is transaminated to glutamate 5 -semialdehyde,
which is oxidized to form glutamate.

536

CHAPTER 17 Amino Acid Metabolism

coo°

coo e

coo°

1 h 2 o

H 3 N— CH
|

H 3 N — CH
|

H 2 0

CH 2 V*

ch 2

1 V

CH 2 urea

ch 2

IPLPI ch 2

h 2 o

ch 2

c=o

©nh 3

c — nh 2

Ornithine

Glutamate 5-semialdehyde

©nh 2 nad©+h 2 o-^

Arginine

NADH + 2H©

COO'

m CH
HN CH 2

w /

HC — CH 2

H 2 0 1/2 o 2

< w

A^Pyrroline 5-carboxylate

coo°

©/ \

h 3 n ch 2

\ /

h 2 c— ch 2

Proline

▲ Figure 17.30

Principal catabolic pathways for arginine,
proline, and histidine.

hUN-

COO'

-CH

ChH 2

COO'

Glutamate

HUN

-N-

| Tetrahydrofolate

5-Formimino-
tetrahydrofolate

A/-Formiminoglutamate Histidine

▲ Proline utilization A flavoprotein. This
enzyme from Bradyrhizobium japonicum
combines the first two enzymes in the
proline degradation pathway into a large
complex consisting of six subunits of bifunc-
tional proteins. The two identical subunits
of one core dimer are colored blue and pur-
ple and the entire structure consists of three
such dimers arranged in a circle. The bound
FAD and NAD© coenzymes are shown as
space-filling models. This enzyme presum-
ably confers a selective advantage over
species containing two separate enzymes
so why hasn’t it evolved in eukaryotes?

[PDB 3HAZ]

Proline is converted to glutamate in three steps. The first step is an oxidation reac-
tion catalyzed by the FAD -containing enzyme proline dehydrogenase. The electron ac-
ceptor is sometimes molecular oxygen although other acceptors can be used. The prod-
uct of the first reaction is A^pyrroline 5-carboxylate (P5C) that exists in equilibrium
with the open-chain form, glutamate 5 -semialdehyde. Glutamate 5 -semialdehyde is
converted to glutamate by the action of NAD® -dependant P5C dehydrogenase. Note
that the conversion of A^pyrroline 5-carboxylate to glutamate 5 -semialdehyde is spon-
taneous as in the proline synthesis pathway (Section 17.3D).

The first two enzymes in this pathway are separate enzymes in all eukaryotes and
most bacteria but in some species of bacteria the two genes for these enzymes have fused
to create a bifunctional hexameric protein that catalyzes both reactions. This is kinetically
advantageous since the intermediates (A^pyrroline 5-carboxylate and glutamate 5-semi-
aldehyde) do not dissociate from the complex before being converted to glutamate.

The major pathway for histidine degradation also produces glutamate. Histidine
undergoes nonoxidative deamination, hydration, and ring opening to form
N-formiminoglutamate. The formimino moiety ( — CH = NH 2 ®) is then trans-
ferred to tetrahydrofolate, forming 5-formiminotetrahydrofolate and glutamate. 5-
Formiminotetrahydrofolate is then enzymatically deaminated to form 5,10-methenylte-
trahydrofolate. The one-carbon (methenyl) group of this tetrahydrofolate derivative
can be used in pathways such as pyrimidine synthesis (Section 18.6).

C. Glycine and Serine

There are two pathways for the breakdown of serine (Figure 17.31). A small amount of
serine is converted directly to pyruvate by the action of serine dehydratase, a PLP-
dependent enzyme. Most serine, however, is converted to glycine by the action of serine
hydroxymethyltransferease. This is the same reaction that results in synthesis of glycine
in the biosynthesis pathway (Figure 17.16) and it is a reaction that produces 5,10-methylene
tetrahydrofolate (5,10-methylene THF).

Some glycine can be converted to serine by the reverse reaction of serine hydroxyl -
methyltransferase and the glycine carbon atoms can end up in pyruvate when the serine
molecules are deaminated. However, the major pathway for degradation of glycine in all
species is conversion to NH 4 ® and HCQ 3 ® by the glycine cleavage system.

17.6 Amino Acid Catabolism

537

Catalysis by the glycine cleavage system requires an enzyme complex containing
four nonidentical subunits. PLP, lipoamide, and FAD are prosthetic groups, and NAD®
and tetrahydrofolate (THF) are cosubstrates. Initially, glycine is decarboxylated and
the — CH 2 — NH 3 ® group is transferred to lipoamide. Then, NH 4 ® is released, and
the remaining one-carbon group is transferred to tetrahydrofolate to form 5,10-
methylenetetrahydro folate (5,10-methylene THF). Reduced lipoamide is oxidized by
FAD and FADH 2 reduces the mobile carrier NAD®.

As shown in Figure 17.32 the glycine cleavage system is another example of a
lipoamide swinging arm mechanism similar in principle to that of pyruvate dehydroge-
nase (Section 13.1). Although glycine breakdown is reversible in vitro , the glycine cleav-
age system catalyzes an irreversible reaction in cells. The irreversibility of the reaction
sequence is due in part to the K m values for the products ammonia and methylene-
tetrahydrofolate that are far greater than the concentrations of these compounds in vivo.

D. Threonine

There are several routes for the degradation of threonine. In the major pathway, threo-
nine is oxidized to 2-amino-3-ketobutyrate in a reaction catalyzed by threonine dehy-
drogenase (Figure 17.33). 2-Amino-3-ketobutyrate can undergo thiolysis to form acetyl
CoA and glycine. Another route for threonine catabolism is cleavage to acetaldehyde
and glycine by the action of threonine aldolase. Threonine aldolase is actually a minor
activity of serine hydroxymethyltransferase in many tissues and organisms. Acetalde-
hyde can be oxidized to acetate by the action of acetaldehyde dehydrogenase and acetate
can be converted to acetyl CoA by acetyl- CoA synthetase.

A third route for threonine catabolism in mammals is deamination to ct-ketobu-
tyrate. This reaction is catalyzed by serine dehydratase, the same enzyme that catalyzes
the conversion of serine to pyruvate. This reaction produces a-ketobutyrate for synthesis
of isoleucine in most species (Section 17.3C). a-Ketobutyrate can be converted to propi-
onyl CoA in the degradative pathway and propionyl CoA is a precursor of the citric acid
cycle intermediate succinyl CoA (Section 16.7 F). Threonine can thus produce either
succinyl CoA or glycine + acetyl CoA depending on the pathway by which it is degraded.

E. The Branched Chain Amino Acids

Leucine, valine, and isoleucine are degraded by related pathways (Figure 17.34). The same
three enzymes catalyze the first three steps in all pathways. The first step, transamination,
is catalyzed by branched chain amino acid transaminase.

The second step in the catabolism of branched chain amino acids is catalyzed by
branched chain ct-keto acid dehydrogenase. In this reaction, the branched chain ct-keto
acids undergo oxidative decarboxylation to form branched chain acyl CoA molecules
one carbon atom shorter than the precursor ct-keto acids. Branched chain ct-keto acid
dehydrogenase is a multienzyme complex containing lipoamide and thiamine pyrophos-
phate (TPP) and requires NAD® and coenzyme A. Its catalytic mechanism is similar to

COO 1

,©

c=o

ch 3

Pyruvate

Serine

dehydratase

(PLP)

•nhP

coo'

,©

H,N— CH

CH ?

Serine

hydroxymethyl

transferase

^-THF

^->5, 10-Methylene THF
^H 2 0

COO

,©

ch 2

©nh 3

Glycine

Glycine-cleavage

system

(PLP, lipoamide,
FAD)

NAD®+ H 2 0

THF

NADH + H®

5,10-Methylene THF

HC0 3 ° + NH 4 ©

▲ Figure 17.31

Catabolism of serine and glycine.

The pathway from propionyl CoA to
succinyl CoA is shown in detail in
Figure 16.22.

◄ Figure 17.32

Glycine cleavage system. A lipoamide swing-
ing arm is attached to the core structural
component (H-protein). The swinging arm
visits the active sites of the three enzymes
of the pathway.

538 CHAPTER 17 Amino Acid Metabolism

coo°

C = 0

T 2

ch 3

a-Ketobutyrate

COO'

,©

NADH + H©

COO'

i©

NH d ©

H,N — CH

Serine

dehydratase

(PLP)

Minor pathway

NAD©

H,N — CH

a-Keto acid
decarboxylase

HS-CoA

■NAD©

NADH

co 2

S-CoA

C = 0

ch 2

ch 3

Propionyl CoA

H C OH Threonine dehydrogenase
Major pathway

CH 3

Threonine

aldolase

Minor

pathway

-> C = 0

ch 3

Acetaldehyde

COO'

h 3 n — ch 2

Glycine

Glycine-cleavage system

C=0

ch 3

2-Amino-3-ketobutyrate

HS-CoA S-CoA

2-Amino-

3-ketobutyrate

lyase

-> C=0

ch 3

Acetyl CoA

coo (

h 3 n — ch 2

Glycine

Glycine-cleavage system

▲ Figure 17.33

Alternate routes for the degradation of threonine.

H,C

©nh 3

CH — CH, — CH — COO

Leucine

H,C.

h 3 c

©nh 3

CH — CH — C00 (

Valine

Transamination

Branched chain
amino acid transaminase

(PLP) Glutamate

- a-Ketoglutarate

HdC — H?C

©NH 3

CH — CH — COO'

3 Isoleucine

Oxidative

decarboxylation

>:-Keto acid

R — C — COO 1

- NAD® + HS-CoA

Branched chain
a-keto acid dehydrogenase

(Lipoamide, TPP) ^NADH + C0 2

Acyl CoA

R — C— S-CoA

Dehydrogenation Acyl-CoA dehydrogenase

ETF:FAD ^n^QH 2
^ETF:FADH 2 Q

yv 2

◄ Figure 17.34

Catabolism of the branched chain amino
acids. R represents the side chain of
leucine, valine, or isoleucine.

H,C— C = CH — C— S-CoA

H ? C = C — C— S-CoA

HoC — CH = C — C— S-CoA

CH,

3 reactions

Acetyl CoA Acetoacetate

CH,

4 reactions

Propionyl CoA

CH,

3 reactions

Acetyl CoA Propionyl CoA

Acetyl CoA

Succinyl CoA

17.6 Amino Acid Catabolism

539

that of the pyruvate dehydrogenase complex (Section 13.1) and the a-ketoglutarate de-
hydrogenase complex (Section 13.3#4), and it contains the same dihydrolipoamide dehy-
drogenase (E 3 ) subunits as those found in the other two dehydrogenase complexes.

Branched chain acyl Co A molecules are oxidized by an FAD -containing acyl- Co A
dehydrogenase in a reaction analogous to the first step in fatty acyl CoA oxidation
(Figure 16.19). The electrons removed in this oxidation step are transferred via the elec-
tron transferring flavoprotein (ETF) to ubiquinone (Q).

At this point, the steps in the catabolism of branched chain amino acids diverge. All
the carbons of leucine are ultimately converted to acetyl CoA, so leucine is purely keto-
genic. Valine is ultimately converted to propionyl CoA. As in the degradation of threo-
nine, propionyl CoA is converted to succinyl CoA that enters the citric acid cycle. Valine
is glucogenic. The isoleucine degradation pathway leads to both propionyl CoA and
acetyl CoA. Isoleucine is therefore both glucogenic (via succinyl CoA formed from pro-
pionyl CoA) and ketogenic (via acetyl CoA). Thus, although the initial steps in the
degradation of the three branched chain amino acids are similar, their carbon skeletons
have different fates — at least in animals.

Remember that the distinction between
ketogenic and glucogenic pathways is
only relevant in animals because all
other species can convert acetyl CoA to
glucose.

F. Methionine

One major role of methionine is conversion to the activated methyl donor S-adenosyl-
methionine (Section 7.3). Transfer of the methyl group from S-adenosylmethionine to
a methyl acceptor leaves S-adenosylhomocysteine that is degraded by hydrolysis to
homocysteine and adenosine (Figure 17.35). Homocysteine can either be methylated by

1 ©

H 3 C— S — CH 2 — CH 2 — CH — coo°
Methionine

H 2 0 + ATP
^ + pPi ^

h 3 c— s — ch 2 — ch 2 — ch— coo©

S-Adenosylmethionine

h 3 c — X ^

©nh 3

1 ©

S — CH 2 — CH 2 — CH — COO°

h 3 c — ch 2 — C— S-CoA
Propionyl CoA

◄ Figure 17.35

Conversion of methionine to cysteine and pro-
pionyl CoA. X in the second step represents
any of a number of methyl-group acceptors.

CO,

NADH

V- NAD©

v HS-CoA

h 3 c — ch 2 — c — coo©

a-Ketobutyrate

°OOC — CH — CH 2 — SH
©NH 3 Cysteine

©ooc — ch — ch 2 — s — ch 2 — ch 2 — ch— coo©

OOC — CH — CH?

HoO

-OH -
Serine

(PLP)

S-Adenosy I homocysteine

h 2 o

Adenosine

©NH 3

1 ©

HS — CH 2 — CH 2 — CH — COO©

Homocysteine

540

CHAPTER 17 Amino Acid Metabolism

coo°

H,N — CH

CH.

Cysteine

^ 0 2

SH©

coo'

I©

H,N — CH

CH,

S0 2 °

Cysteinesulfinate

a- Keto-
^glutarate

^Glutamate

(PLP)

coo 0

C =0

ch 2

so 2

/3-Sulfinylpyruvate

Nonenzymatic

desulfurylation

-H®

k S0 2

coo°

C =0

ch 3

Pyruvate

▲ Figure 17.36

Conversion of cysteine to pyruvate.

BOX 17.5 PHENYLKETONURIA IS A DEFECT IN TYROSINE
FORMATION

One of the most common disorders of
amino acid metabolism is phenylke-
tonuria (PKU). The disease is caused
by a mutation in the gene that encodes
phenylalanine hydroxylase ( PAH
gene on chromosome 12q: OMIN
MIN=261600). Affected individuals
are unable to convert dietary pheny-
lalanine to tyrosine so the blood of
children with this disease contains very
high levels of phenylalanine and low
levels of tyrosine. Instead of being con-
verted to tyrosine, phenylalanine is
metabolized to phenylpyruvate in the
reverse of the transamination reaction
shown in Figure 17.20. (Transamina-
tion of phenylalanine does not occur in
unaffected individuals because the K m
of the transaminase for phenylalanine
is much higher than the normal con-
centration of phenylalanine.) Elevated
levels of phenylpyruvate and its deriva-
tives inhibit brain development.

Newborns are routinely screened
for PKU by testing for elevated levels of
phenylpyruvate in the urine or of
phenylalanine in the blood during the
first days after birth. Phenylalanine
hydroxylase-deficient individuals often
develop normally if the dietary intake of
phenylalanine is strictly limited during
the first decade of life. Some women
with PKU must restrict their dietary in-
take of phenylalanine during pregnancy

to ensure proper fetal development. Ele-
vated levels of phenylalanine are also
observed in individuals with deficiencies
in dihydropteridine reductase or 4a-
carbinolamine dehydratase or defects in
the biosynthesis of tetrahydrobiopterin
because each of these disorders results in
impairment of the hydroxylation of
phenylalanine.

Control of diet can successfully
treat PKU but the restrictions exclude
many natural, protein-rich foods such
as meat, fish, milk, bread, and cake. The
food of this strict diet is not appetizing.
Tests have been performed by feeding
PKU victims an enzyme that catalyzes
degradation of phenylalanine to ammo-
nia and a nontoxic carbon product. This
enzyme does not fully replace dietary
restriction of phenylalanine but it may
increase a patient’s tolerance for pro-
tein-containing foods.

▲ Newborn infants are tested for phenylke-
tonuria by analyzing blood drawn from the
heel of the foot.

5-methyltetrahydrofolate to form methionine or it can react with serine to form cys-
tathionine that can be cleaved to cysteine and cr-ketobutyrate. We encountered this se-
ries of reactions earlier as part of a pathway for the formation of cysteine (Figure 17.18).
By this pathway, mammals can form cysteine using a sulfur atom from the essential
amino acid methionine. a-Ketobutyrate is converted to propionyl CoA by the action of
an cr-keto acid dehydrogenase. Propionyl CoA can be further metabolized to succinyl
CoA, so methionine is glucogenic.

G. Cysteine

The major route of cysteine catabolism is a three-step pathway leading to pyruvate
(Figure 17.36). Therefore, cysteine is glucogenic. Cysteine is first oxidized to cysteinesul-
finate that loses its amino group by transamination to form /3-sulfinylpyruvate. Nonen-
zymatic desulfurylation produces pyruvate.

17.6 Amino Acid Catabolism 541

coo 0

e I

H 3 N — C — H

H
H

Phenylalanine

COO G

H 3 N — C — H

H
H

Tyrosine

H ? N

3 reactions

CH — CH — CHq

OH OH

5,6,7,8-Tetrahydrobiopterin

OH OH
4u-Carbinolamine

C =

/ 5

ICOO'

,©

= c

6 \

OOC4 H

Fumarate
+

H,C-

CH-> — coo

I©

Acetoacetate

4a-Carbinolamine

dehydratase

■» h 2 o

OH OH

Dihydrobiopterin
(Quinonoid form)

▲ Figure 17.37

Conversion of phenylalanine and tyrosine to fumarate and acetoacetate. The tetrahydrobiopterin
cofactor is regenerated via dehydration and NADH-dependent reduction.

H. Phenylalanine, Tryptophan, and Tyrosine

The aromatic amino acids share a common pattern of catabolism. In general, the path-
ways begin with oxidation, followed by removal of nitrogen by transamination or hy-
drolysis and then ring opening coupled with oxidation.

The conversion of phenylalanine to tyrosine, catalyzed by phenylalanine hydroxy-
lase, is an important step in the catabolism of phenylalanine (Figure 17.37). It also
serves as a source of tyrosine in animals since they lack the normal chorismate pathway
for tyrosine synthesis. The phenylalanine hydroxylase reaction requires molecular oxy-
gen and the reducing agent tetrahydrobiopterin. One oxygen atom from 0 2 is incorpo-
rated into tyrosine and the other is converted to water.

Tetrahydrobiopterin is regenerated in two steps. 4a-Carbinolamine dehydratase
catalyzes the dehydration of the first oxidized product and prevents its isomerization to
an inactive form in which the side chain is on C-7, not C-6. Dihydropteridine reductase
catalyzes the reduction of the resulting quinonoid dihydrobiopterin to 5,6,7,8-tetrahy-
drobiopterin in a reaction that requires NADH. Tetrahydrobiopterin is also a reducing
agent in the biosynthesis of nitric oxide from arginine (Section 17.4C).

542 CHAPTER 17 Amino Acid Metabolism

®NH 3

2 — CH— COO 0
Tryptophan

8 reactions

©NH 3

G OOC- (CH 2 ) 3 — c - coo°
u-Ketoadipate

6 reactions

-COO'
Alanine

2 H 3 C — C — S-CoA + 2 C0 2
2 Acetyl CoA

©NH 3

H 2 N — (CH 2 ) 4 — CH — COO 0
Lysine

u-Ketoglutarate

H 2 0

NADPH + H 0
NADP®

COO'

i©

©NH,

CH — N — (CH 2 ) 4 — CH — COO 1
I H
CH 2

CH 2 Saccharopine

COO'

h 2 o

Glutamate -

NAD 0
NADH + H 0

©NH,

C — (CH 2 ) 3 — CH — COO 0

u-Ami noadipate 5-semialdehyde

H 2 0

NADP 0

Oxidation |\|ADPH + 2H®

\ t

©NH 3

°OOC— (CH 2 ) 3 — CH — COO 0
u-Ami noadipate

u-Ketoglutarate

Transamination

Glutamate

o 11 o

°ooc— (ch 2 ) 3 — c — coo°

u-Ketoadipate
6 reactions

2 Acetyl CoA + 2 C0 2
▲ Figure 17.39

Conversion of lysine to acetyl CoA.

Further degradation of uric acid is
described in Section 18.8.

▲ Figure 17.38

Conversion of tryptophan to alanine and acetyl CoA.

The catabolism of tyrosine begins with the removal of its a-amino group in a
transamination reaction with a-ketoglutarate. Subsequent oxidation steps lead to ring
opening and eventually to the final products, fumarate and acetoacetate. This fumarate
is cytosolic and is converted to glucose. Acetoacetate is a ketone body. Thus, tyrosine is
both glucogenic and ketogenic.

The indole ring system of tryptophan has a more complex degradation pathway
that includes two ring- opening reactions. The major route of tryptophan catabolism in
the liver and many microorganisms leads to a-ketoadipate and ultimately to acetyl CoA
(Figure 17.38). Alanine, produced early in tryptophan catabolism, is transaminated to
pyruvate. Thus, the catabolism of tryptophan is both ketogenic and glucogenic.

I. Lysine

The main pathway for the degradation of lysine generates the intermediate saccharopine,
the product of the condensation of a-ketoglutarate with lysine (Figure 17.39). Sequential
oxidation reactions produce u- amino adipate that loses its amino group by transamination
with o'-ketoglutarate to become a-ketoadipate. a-Ketoadipate is subsequently converted to
acetyl CoA by the same steps that occur in the degradation of tryptophan. Like leucine,
lysine is ketogenic (these two are the only common amino acids that are purely ketogenic).

17.7 The Urea Cycle Converts Ammonia
into Urea

High concentrations of ammonia are toxic to cells. Different organisms have evolved
different strategies for eliminating waste ammonia. The nature of the excretory product
depends on the availability of water. In many aquatic organisms, ammonia diffuses di-
rectly across the cell membranes and is diluted by the surrounding water. This route is
inefficient in large terrestrial multicellular organisms and the buildup of ammonia in-
side internal cells must be avoided.

Most terrestrial vertebrates convert waste ammonia to urea, a less toxic product
(Figure 17.40). Urea is an uncharged and highly water-soluble compound produced in
the liver and carried in the blood to the kidneys where it is excreted as the major solute
of urine. (Urea was first described around 1720 as the essential salt of urine. The name
“urea” is derived from “urine”) Birds and many terrestrial reptiles convert surplus am-
monia to uric acid, a relatively insoluble compound that precipitates from aqueous
solution to form a semisolid slurry. Uric acid is also a product of the degradation of
purine nucleotides by birds, some reptiles, and primates.

The synthesis of urea occurs almost exclusively in the liver. Urea is the product of a
set of reactions called the urea cycle — a pathway discovered by Hans Krebs and Kurt
Henseleit in 1932 several years before Krebs discovered the citric acid cycle. Several ob-
servations led to the identification of the urea cycle; for example, slices of rat liver can
bring about the net conversion of ammonia to urea. Synthesis of urea by these prepara-
tions is markedly stimulated when the amino acid ornithine is added and the amount of
urea synthesized greatly exceeds the amount of ornithine that is added, suggesting that
ornithine acts catalytically. Finally, it was known that high levels of the enzyme arginase
occur in the livers of all organisms that synthesize urea.

17.7 The Urea Cycle Converts Ammonia into Urea 543

H,N

C = 0

H 2 N

Urea

Uric acid

◄ Figure 17.40
Urea and uric acid.

O— ADP

HO
Bicarbonate

First ATP

ADP

A. Synthesis of Carbamoyl Phosphate

The ammonia released by oxidative deamination of glutamate reacts with bicarbonate
to form carbamoyl phosphate. This reaction requires two molecules of ATP and is
catalyzed by carbamoyl phosphate synthetase (Figure 17.41). This enzyme is present in
all species since carbamoyl phosphate is an essential precursor in pyrimidine biosynthe-
sis and it’s also required in the synthesis of arginine in species that don’t have a urea
cycle. Mammals have two versions of this enzyme. The cytosolic version is called car-
bamoyl phosphate synthetase II and it uses glutamine rather than ammonia as the
nitrogen donor. This is the enzyme used in pyrimidine synthesis (Section 18.3). The
bacterial enzymes also use glutamine. The second mammalian version, carbamoyl
phosphate I, is the one involved in the urea cycle. It is one of the most abundant en-
zymes in liver mitochondria accounting for as much as 20% of the protein of the mito-
chondrial matrix. The nitrogen atom of carbamoyl phosphate is incorporated into urea
via the urea cycle.

Ammonia

,0.

h 2 n— c— o— P— o°

HO O 0

Tetrahedral intermediate

B. The Reactions of the Urea Cycle

The first nitrogen atom of urea is contributed by carbamoyl phosphate and the second
is derived from aspartate. The synthesis of urea takes place while the intermediates are
covalently bound to an ornithine skeleton. Ornithine is regenerated when urea is re-
leased and it re-enters the urea cycle. Thus, ornithine acts catalytically in the synthesis of
urea (Figure 17.42). The carbon, nitrogen, and oxygen atoms of ornithine are not

H 2 I\ — C —

Urea

H 2 N — C

x o1h

Carbamate

°0— P— O— ADP

A®

Second ATP

^ADP

h 2 n — c — o— P — o

o 0

Carbamoyl phosphate

▲ Figure 17.41

Synthesis of carbamoyl phosphate catalyzed by
carbamoyl phosphate synthetase I. The reac-
tion involves two phosphoryl-group transfers.
First, nucleophilic attack by bicarbonate on
ATP produces carboxy phosphate and ADP.
Next, ammonia reacts with carboxy phos-
phate, forming a tetrahedral intermediate.
Elimination of a phosphate group produces
carbamate. A second phosphoryl-group
transfer from another ATP forms carbamoyl
phosphate and ADP. Structures in brackets
remain enzyme bound during the reaction.

▲ Figure 17.42

The urea cycle. The blue rectangular box represents ornithine.

544 CHAPTER 17 Amino Acid Metabolism

BOX 17.6 DISEASES OF AMINO ACID METABOLISM

Hundreds of human metabolic dis-
eases involving single-gene defects
(often termed inborn errors of metab-
olism) have been discovered. Many are
due to defects in the breakdown of
amino acids. We have already discussed
phenylketonuria, the defect in tyrosine
formation from phenylalanine (Box 17.5).
A few more examples are mentioned
here. Defects in some pathways are se-
vere and even life-threatening; defects
in other pathways can result in less se-
vere symptoms. The results indicate
that some amino acid degradation
pathways are almost dispensable whereas
others are essential for survival follow-
ing birth.

Alkaptonuria

The first metabolic disease to be char-
acterized as a genetic defect was alkap-
tonuria, a rare disease in which one of
the intermediates in the catabolism of
phenylalanine and tyrosine (homogen-
tisate) accumulates (Figure 17.37). A
deficiency of homogentisate dioxyge-
nase, the enzyme that catalyzes oxida-
tive cleavage of this intermediate, pre-
vents further metabolism of this
catabolite. The gene is HGD on chro-
mosome 3 (OMIM MIM=203500).

Solutions of homogentisate turn dark
on standing because this compound is
converted to a pigment. Alkaptonuria
was recognized by observing the dark-
ening of urine. Individuals with alkap-
tonuria are prone to develop arthritis,
but it is not known how the metabolic
defect produces this complication; pos-
sibly it is from the deposit of pigments
in bones and connective tissues.

Cystinuria

If there is a defect in kidney transport
of cysteine and the basic amino acids,
then cysteine accumulates in blood and
oxidizes to cystine producing a condition
called cystinuria. Cystine has a low sol-
ubility and forms calculi. Patients suf-
fering from cystinuria drink large
amounts of water to dissolve these
stones or are given compounds that
react with cystine to form soluble de-
rivatives. (See OMIM MIM=220100.)

Gyrate Atrophy

A defect in ornithine transaminase ac-
tivity causes the metabolic disease gyrate
atrophy of the choroid and retina of the
eyes. The affected gene is OAT on chro-
mosome 10 (OMIM MIM=258870).
Gyrate atrophy leads to tunnel vision

and later to blindness. The progress of
this disorder can be slowed by restricting
the dietary intake of arginine or by the
administration of pyridoxine.

Maple Syrup Urine Disease

Patients suffering from maple syrup
urine disease excrete urine that smells
like maple syrup. The disease is caused
by a genetic defect at the second step in
catabolism of branched chain amino
acids — the step catalyzed by the
branched chain a-keto acid dehydroge-
nase complex. Those afflicted with this
disease have short lives unless they fol-
low a diet very low in branched chain
amino acids. (OMIM MIM=248600)

Nonketotic Hyperglycinemia
(Glycine Encephalopathy)

Defects in the enzyme complex that
catalyzes glycine cleavage lead to the ac-
cumulation of large amounts of glycine
in body fluids. This is the main bio-
chemical symptom of a disease called
nonketotic hyperglycinemia. Most indi-
viduals with this disorder have severe
mental deficiencies and die in infancy.
The severity of the disease indicates the
crucial importance of the glycine cleav-
age system. (OMIM MIM=605899)

KEY CONCEPT

All species need to eliminate ammonia
produced by degradation reactions. Some
can excrete it directly while others have
to convert it to less toxic compounds that
are subsequently excreted.

exchanged in the urea cycle. Its role as a catalyst is more obvious than the role of ox-
aloacetate in the citric acid cycle (Section 13.3) but the principle is the same.

The actual urea cycle reactions are more complex than the simple scheme shown in
Figure 17.42. This is because the first reaction occurs in the mitochondrial matrix and
the other three occur in the cytosol (Figure 17.43). Two transport proteins connecting
the mitochondrial matrix and the cytosol are required: the citrulline-ornithine ex-
changer and the glutamate-aspartate translocase.

1. The cycle begins when carbamoyl phosphate reacts in the mitochondrion with or-
nithine to form citrulline in a reaction catalyzed by ornithine transcarbamoylase.
This step incorporates the nitrogen atom originating from ammonia into cit-
rulline; citrulline thus contains half the nitrogen destined for urea. Citrulline is
then transported out of the mitochondrion in exchange for cytosolic ornithine.

2. The second nitrogen atom destined for urea comes from aspartate and is incorpo-
rated when citrulline condenses with aspartate to form argininosuccinate in the cy-
tosol. This ATP-dependent reaction is catalyzed by argininosuccinate synthetase.
Most aspartate in cells originates in mitochondria although aspartate is sometimes
generated in the cytosol. Mitochondrial aspartate enters the cytosol in exchange for
cytosolic glutamate. (This translocase reaction is part of the malate-aspartate shut-
tle we described in Section 14.12.)

3. Argininosuccinate is cleaved nonhydrolytically to form arginine plus fumarate in
an elimination reaction catalyzed by argininosuccinate lyase. Arginine is the immediate

17.7 The Urea Cycle Converts Ammonia into Urea 545

Figure 17.43 ►
Urea cycle.

NH 2

r° —

0P0 3 ©

Carbamoyl phosphate

2 ADP + P:

2 ATP HCO

,©

Carbamoyl phosphate synthetase I

NHo

NAD(P)H + 2 H®<

Glutamate dehydrogenase

NAD(P)® + H 2 0

MITOCHONDRIAL

MATRIX

©NH 3

Ornithine

©NH 3

Citrulline

H,N-

Aspartate

CYTOSOL

c = o

nh 2

Urea

H,0

Ornithine -

Arginase (4)

©NH 2

c — nh 2

ChH 2

C H 2

CH — COO®

©nh 3

Arginine

Citrulline

©NH,

Aspartate

ATP

AMP + PPi

COO'

,©

Argininosuccinate lyase

,©

coo'

COO'

,©

c — r — ch
I H |

NH CH 2

I I

ch 2 coo'

ch 2

CH — COO®

©nh 3

Argininosuccinate

COO'

c = o

ch 2

CH,

COO®

a-Ketoglutarate

coo®

h 3 n —

ch 2

CH 2

coo®

Glutamate

Fumarate

precursor of urea. (Together, the second and third steps of the urea cycle exem-
plify a strategy for donating the amino group of aspartate. We will encounter this
strategy twice more in the next chapter as part of purine biosynthesis. The key
processes are a nucleoside triphosphate-dependent condensation, followed by the
elimination of fumarate.)

4. Finally, the guanidinium group of arginine is hydrolytically cleaved to form or-
nithine and urea in a reaction catalyzed by arginase. Arginase has a pair of Mn®
ions in its active site and this binuclear manganese cluster binds a molecule of
water forming a nucleophilic hydroxide ion that attacks the guanidinium carbon
atom of arginine. The ornithine generated by the action of arginase is transported

546

CHAPTER 17 Amino Acid Metabolism

into the mitochondrion where it reacts with carbamoyl phosphate to support con-
tinued operation of the urea cycle.

The overall reaction for urea synthesis is

NH 3 + HCO 3 0 + Aspartate + 3 ATP *

Urea + Fumarate + 2 ADP + 2 Pj + AMP + PPj (17.5)

The two nitrogen atoms of urea are derived from ammonia and aspartate. The carbon
atom of urea comes from bicarbonate. Four equivalents of ATP are consumed per mol-
ecule of urea synthesized. Three molecules of ATP are converted to two ADP and one
AMP during the formation of one molecule of urea and the hydrolysis of inorganic py-
rophosphate accounts for cleavage of the fourth phosphoanhydride bond.

The carbon skeleton of fumarate is converted to glucose and C0 2 . Cytosolic fu-
marate does not enter the citric acid cycle (which occurs in mitochondria) but instead is
hydrated to malate by the action of a cytosolic fumarase. Malate is oxidized to oxaloac-
etate by the action of malate dehydrogenase and oxaloacetate enters the pathway of glu-
coneogenesis. This fate is shared by the fumarate produced during tyrosine degradation
(Section 17.6H).

C. Ancillary Reactions of the Urea Cycle

The reactions of the urea cycle convert equal amounts of nitrogen from ammonia and from
aspartate into urea. Many amino acids can function as amino -group donors via transami-
nation reactions with a-ketoglutarate to form glutamate. Glutamate can undergo either
transamination with oxaloacetate to form aspartate or deamination to form ammonia.
Both glutamate dehydrogenase and aspartate transaminase are abundant in liver mitochon-
dria and catalyze near-equilibrium reactions. The concentrations of ammonia and aspar-
tate must be approximately equal for efficient synthesis of urea and elimination of nitrogen.

Consider the theoretical case of a relative surplus of ammonia (Figure 17.44a). In
this situation, the near-equilibrium reaction catalyzed by glutamate dehydrogenase
would proceed in the direction of glutamate formation. Elevated concentrations of glu-
tamate would then result in increased flux to aspartate through aspartate transaminase,
also a near- equilibrium step. In contrast, when excess aspartate is present the net flux in
the reactions catalyzed by glutamate dehydrogenase and aspartate transaminase would
occur in the opposite direction to provide ammonia for urea formation (Figure 17.44b).

Figure 17.44 ►

Balancing the supply of nitrogen for the urea
cycle. Two theoretical situations are de-
scribed: (a) NH 3 in extreme excess and
(b) aspartate in extreme excess. Flux
through the near-equilibrium reactions cat-
alyzed by glutamate dehydrogenase and as-
partate transaminase reverses, depending on
the relative amounts of ammonia and amino
acids.

(a) NH 3 in excess

(b) Aspartate in excess

NH 3 «

u-Ketoglutarate

NADH

NAD

Oxaloacetate Aspartate

Carbamoyl _ Citrulline ^

phosphate *

Urea <-

Urea

cycle

Urea «■

Urea

cycle

17.8 Renal Glutamine Metabolism Produces Bicarbonate

547

MUSCLE

◄ Figure 17.45
Glucose-alanine cycle.

Some amino acids are deaminated in muscle, not in the liver. Glycolysis — a major
source of energy in muscle — produces pyruvate. The transfer of amino groups from
a - amino acids to pyruvate generates large amounts of alanine. Alanine travels through the
bloodstream to the liver where it is deaminated back to pyruvate. The amino group is
used for urea synthesis and the pyruvate is converted to glucose by gluconeogenesis. Re-
call that neither of these pathways operates in muscle. Glucose can return to the muscle
tissue. Alternatively, pyruvate can be converted to oxaloacetate that becomes the carbon
chain of aspartate — the metabolite that donates one of the nitrogen atoms of urea. The
exchange of glucose and alanine between muscle and liver is called the glucose-alanine
cycle (Figure 17.45) and it provides an indirect means for muscle to eliminate nitrogen
and replenish its energy supply.

17.8 Renal Glutamine Metabolism
Produces Bicarbonate

The body often produces acids as metabolic end products. The resulting anions are
eliminated in the urine and the protons remain in the body. One example is /3-hydroxy-
butyric acid, a ketone body that is produced in massive amounts during uncontrolled
diabetes mellitus. Another example is sulfuric acid produced during catabolism of the
sulfur- containing amino acids cysteine and methionine. These acid metabolites dissoci-
ate to give protons and the corresponding anion, /3-hydroxybutyrate or sulfate (S0 4 ©).
The blood has an effective buffer system for the protons — they react with bicarbonate
to produce C0 2 that is eliminated by the lungs and H 2 0 (Figure 17.46). While this
system effectively neutralizes the excess hydrogen ions it does so at the cost of depleting
blood bicarbonate. Bicarbonate is replenished by glutamine catabolism in the kidneys.

In the kidneys, the two nitrogen atoms of glutamine are removed by the sequential
action of glutaminase and of glutamate dehydrogenase to produce a-ketoglutarate©
and 2 NH 4 ®.

Glutamine > > a-Ketoglutarate® + 2 NH 4 ® (17.6)

Two molecules of the divalent anion a-ketoglutarate can be converted to one molecule
of neutral glucose and four molecules of bicarbonate. The a-ketoglutarate is con-
verted to glucose by oxidation to oxaloacetate, leading to gluconeogenesis. The overall
process (ignoring ATP involvement) is

2 C5H 10 N 2 O3 + 3 0 2 + 6 H 2 0 >

Glutamine

Glucose

The NH 4 ® is excreted in the urine and the HC0 3 ® is added to the venous blood, replacing
the bicarbonate lost in buffering metabolic acids. The excreted NH 4 © is accompanied in
the urine by the anion (e.g., /3-hydroxybutyrate or sulfate) of the original acid metabolite.

Blood

Lungs

▲ Figure 17.46

548 CHAPTER 17 Amino Acid Metabolism

Summary

1 . Nitrogen is fixed in only a few species of bacteria by the nitrogenase-
catalyzed reduction of atmospheric N 2 to ammonia. Plants and
microorganisms can reduce nitrate and nitrite to ammonia.

2. Ammonia is assimilated into metabolites by the reductive anima-
tion of a-ketoglutarate to glutamate, catalyzed by glutamate de-
hydrogenase. Glutamine, a nitrogen donor in many biosynthetic
reactions, is formed from glutamate and ammonia by the action
of glutamine synthetase.

3. The amino group of glutamate can be transferred to an cr-keto
acid in a reversible transamination reaction to form u-ketoglutarate
and the corresponding a - amino acid.

4. Pathways for the biosynthesis of the carbon skeletons of amino
acids begin with simple metabolic precursors such as pyruvate
and citric acid cycle intermediates.

5. In addition to their role in protein synthesis, amino acids serve as
precursors in a number of other metabolic pathways.

6. Protein molecules in all living cells are continually synthesized
and degraded.

7. Amino acids obtained from protein degradation or directly from
food can be catabolized. Catabolism often begins with deamina-
tion, followed by modification of the remaining carbon chains for
entry into the central pathways of carbon metabolism.

8. The pathways for the degradation of amino acids lead to pyru-
vate, acetyl CoA, or intermediates of the citric acid cycle. Amino
acids that are degraded to citric acid cycle intermediates are
glucogenic. Those that form acetyl CoA are ketogenic.

9. Most nitrogen in mammals is excreted as urea that is formed by
the urea cycle in the liver. The carbon atom of urea is derived
from bicarbonate. One amino group is derived from ammonia
and the other from aspartate.

10. The metabolism of glutamine in the kidneys produces the bicar-
bonate needed to neutralize acids produced in the body.

Problems

1. The heterocysts of cyanobacteria contain high concentrations of
nitrogenase. These cells have retained photosystem I (PSI) but
they do not contain photosystem II (PSII). Why?

2. Write the net equation for converting one molecule of a-ketoglu-
tarate into one molecule of glutamine by assimilating two mole-
cules of ammonia in the following coupled reactions: (a) gluta-
mate dehydrogenase and glutamine synthetase and (b) glutamine
synthetase and glutamate synthase. Compare the energy require-
ments of the two pathways and account for any difference.

3. When 15 AT-labeled aspartate is fed to animals the 15 N label quickly
appears in many amino acids. Explain this observation.

4. (a) Three of the 20 common amino acids are synthesized by sim-

ple transamination of carbohydrate metabolites. Write the
equations for these three transamination reactions.

(b) One amino acid can be synthesized by reductive amination of a
carbohydrate metabolite. Write the equation for this reaction.

5. Animals rely on plants or microorganisms for the incorporation
of sulfur into amino acids and their derivatives. However, methio-
nine is an essential amino acid in animals while cysteine is not. If
the donor of a sulfur atom in the conversion of homoserine to
homocysteine by plants is cysteine, outline the overall path by
which sulfur is incorporated into cysteine and methionine in
plants and into cysteine in animals.

6. Serine is a source of one-carbon fragments for certain biosyn-
thetic pathways.

(a) Write the equations that show how two carbon atoms from
serine are made available for biosynthesis.

(b) Assuming that the precursor of serine is produced by glycolysis,
which carbon atoms of glucose are the ultimate precursors of
these one- carbon fragments?

7. Indicate where the label appears in the product for each of the fol-
lowing precursor-product pairs:

(a) 3- [ 14 C] -Oxaloacetate — > Threonine

(b) 3-[ 14 C]-Phosphoglycerate — > Tryptophan

— c— coo 0

Chorismate

Phenylalanine

8. (a) PPT (phosphinothricin) is a herbicide that is relatively safe
for animals because it is not transported from the blood into
the brain and it is rapidly cleared by animal kidneys. PPT ef-
fectively inhibits an enzyme in plant amino acid metabolism
because it is an analog of the amino acid substrate. What
amino acid does PPT resemble?

11 o

h 3 c— p— ch 2 ch 2 ch —coo 0

O© NH 3 ©

PPT

(b) The herbicide aminotriazole inhibits imidazole glycerol
phosphate dehydrogenase. What amino acid pathway is in-
hibited in plants?

NH 2

Aminotriazole

9. Children with phenylketonuria should not consume the artificial
sweetener aspartame (Figure 3.10). Why?

Selected Readings 549

10 . (a) A deficiency of a-keto acid dehydrogenase is the most com-

mon enzyme abnormality in branched chain amino acid ca-
tabolism. Individuals with this disease excrete branched
chain cr-keto acids. Write the structures of the a-keto acids
that would result during the catabolism of leucine, valine,
and isoleucine when this enzyme is defective.

(b) A disorder of amino acid catabolism results in the accumula-
tion and excretion of saccharopine. What amino acid path-
way is involved and what enzyme is defective?

(c) Citrullinemia is characterized by accumulation of citrulline in
the blood and its excretion in the urine. What metabolic path-
way is involved and what enzyme is deficient for this disease?

11. Which amino acids yield the following a-keto acids by transami-
nation?

(a) O

°ooc— c— ch 3

°OOC— CH

(b) O

°ooc— c— ch 2

(d) O

°ooc— c— ch 2

coo 0

12 . Animal muscles use two mechanisms to eliminate excess nitrogen
generated during the deamination of amino acids. What are the
two pathways and why are they necessary?

13. Thiocitrulline and S-methylthiocitrulline prevent experimentally
induced blood vessel dilation, reduced blood pressure, and shock
in animals. What enzyme that produces a gaseous blood vessel di-
lating messenger is being inhibited? Suggest why these two mole-
cules might act as inhibitors of this enzyme.

14 . Why are there so few metabolic diseases associated with defects in
amino acid biosynthesis?

15 . Pathways for the biosynthesis of the 21st, 22nd, and 23rd amino
acids (Section 3.3) are not described in this chapter. Why not? What
are the immediate precursors of three additional amino acids?

16 . The cost of making amino acids, in ATP equivalents, can be calcu-
lated using values for the cost of making each of the precursors
plus the cost of each reaction in the amino acid biosynthesis path-
way. Assuming that the cost of making glyceraldehyde-3-phosphate
is 24 ATP equivalents (Section 15.4C), calculate the cost of mak-
ing serine (Figure 17.15) and alanine (Figure 17.12). How do
your values compare to those in Box 17.3?

Selected Readings

Nitrogen Cycle

Dixon, R., and Kahn, D. (2004). Genetic regulation
of biological nitrogen fixation. Nat. Rev. Microbiol.
2:621-631.

Moisander, P. H., Beinart, R. A., Hewson, I., White,
A. E., Johnson, K. S., Carlson, C. A., Montoya, J. P.,
and Zehr, J. P. (2010). Unicellular cyanobacterial
distributions broaden the oceanic N 2 fixation do-
main. Science 327:1512-1524.

Montoya, J. P., Holl, C. M., Zehr, J. P., Hansen, A.,
Villareal, T. A., and Capone, D. G. (2004). High
rates of N 2 fixation in the oligotrophic Pacific
ocean. Nature 430: 1027-103 1 .

Schimpl, J., Petrilli, H. M., and Blochl, P. E. (2003).
Nitrogen binding to the FeMo-cofactor of nitroge-
nas e.J.Am. Chem. Soc. 125:15772-15778.

Seefeldt, L. C., Hoffman, B. M., and Dean, D. R.
(2009). Mechanism of Mo- dependent nitrogenase.
Annu. Rev. Biochem. 78:701-722.

Amino Acid Metabolism

Fitzpatrick, P. F. (1999). Tetrahydropterin- dependent
amino acid hydroxylases. Annu. Rev. Biochem.
68:355-381.

Haussinger, D. (1998). Hepatic glutamine trans-
port and metabolism. Adv. Enzymol. Relat. Areas
Mol. Biol. 72:43-86.

Huang. X., Holden, H. M., and Raushel, F. M.
(2001). Channeling of substrates and intermedi-
ates in enzyme- catalyzed reactions. Annu. Rev.
Biochem. 70:149-180.

Katagiri, M., and Nakamura, M. (2003). Reappraisal
of the 20th-century version of amino acid metabo-
lism. Biochem. Biophys, Res , Comm. 312:205-208.

Levy, H. L. (1999). Phenylketonuria: old disease,
new approach to treatment. Proc. Natl. Acad. Sci.
USA 96:1811-1813.

Perham, R. N. (2000). Swinging arms and swinging
domains in multifunctional enzymes: catalytic ma-
chines for multistep reactions. Annu. Rev. Biochem.
69:961-1004.

Purich, D. L. (1998). Advances in the enzymology
of glutamine synthesis. Adv. Enzymol. Relat. Areas
Mol. Biol. 72:9-42.

Raushel, F. M., Thoden, J. B., and Holden, H. M.
(2003). Enzymes with molecular tunnels. Acc.
Chem. Res. 36:539-548.

Richards, N. G. and Kilberg, M. S. (2006).
Asparagine synthetase chemotherapy. Annu. Rev.
Biochem. 75:629-654.

Scapin, G., and Blanchard, J. S. (1998). Enzymol-
ogy of bacterial lysine biosynthesis. Adv. Enzymol.
Relat. Areas Mol. Biol. 72:279-324.

Scriver, C. R., Beaudet, A. L., Sly, W. S., and Valle,
D., eds. (1995). The Metabolic Basis of Inherited
Disease,^ ols. 1, 2, and 3. (New York: McGraw-
Hill).

Srivastava, D., Schuermann, J. P., White, T. A.,
Krishnan, N., Sanyal, N., Hura, G. L., Tan, A.,
Henzl, M. T., Becker, D. F., and Tanner, J. J. (2010).
Crystal structure of the bifunctional proline uti-
lization A flavoenzyme from Bradyrhizobium
japonicum. Proc. Natl. Acad. Sci. USA
107:2878-2883.

Wu, G., and Morris, S. M., Jr. (1998). Arginine
metabolism: nitric oxide and beyond. Biochem. J.
336:1-1 7.

Zalkin, H., and Smith, J. L. (1998). Enzymes utiliz-
ing glutamine as an amide donor. Adv. Enzymol.
Relat. Areas Mol. Biol. 72:87-144.

Nucleotide Metabolism

W e have encountered nucleotides and their constituents throughout this
book. Nucleotides are probably best known as the building blocks of DNA
and RNA; however, as we have seen, they are involved in almost all the ac-
tivities of the cell either alone or in combination with other molecules. Some nu-
cleotides (such as ATP) function as cosubstrates, and others (such as cyclic AMP and
GTP) are regulatory compounds.

One of the components of every nucleotide is a purine or pyrimidine base. The
other components are a five-carbon sugar — ribose or deoxyribose — and one or more
phosphoryl groups. The standard bases (adenine, guanine, cytosine, thymine, uracil)
are almost always found as constituents of nucleotides and polynucleotides. All organ-
isms and cells can synthesize purine and pyrimidine nucleotides because these mole-
cules are essential for information flow. In non- dividing cells, nucleotide biosynthesis is
almost exclusively devoted to the production of ribonucleotides for RNA synthesis and
various nucleotide cofactors. Deoxyribonucleotides are required for DNA replication in
dividing cells and consequently, deoxynucleotide synthesis is closely linked to cell divi-
sion. Its study is particularly important in modern medicine since synthetic agents that
inhibit deoxyribonucleotide synthesis are useful as therapeutic agents against cancer.

We begin this chapter with a description of the biosynthesis of purine and pyrimi-
dine nucleotides. Next, we present the conversion of purine and pyrimidine ribonu-
cleotides to their 2'-deoxy forms, the forms incorporated into DNA. We then discuss
how purines and pyrimidines obtained from the breakdown of nucleic acids or extracel-
lularly from food can be incorporated directly into nucleotides — a process called salvage.
The salvage pathways conserve energy by recycling the products of nucleic acid turnover.
Finally, we examine the biological degradation of nucleotides. The breakdown of purines
leads to the formation of potentially toxic compounds that are excreted, whereas the
breakdown of pyrimidines leads to readily metabolized products.

18.1 Synthesis of Purine Nucleotides

The identification of the enzymes and intermediates in the pathway for the synthesis
of the two purine nucleotides, adenosine 5 '-monophosphate (AMP) and guanosine

[Sven] Furberg, reasoning with
marked brilliance and luck from
data that were meagre but
included his own x-ray studies,
got right the absolute three-
dimensional configuration of the
individual nucleotide . . . " Furberg 's
nucleotide . . . was absolutely es-
sential to us. " Crick told me.

Horace Freeland Judson (1996), The
Eighth Day of Creation, p. 94.

Top: Methotrexate, one of the most commonly used anticancer drugs. Methotrexate is an analog of folate that inhibits the
reaction cycle generating deoxythymidylate for DNA synthesis.

550

18.1 Synthesis of Purine Nucleotides 551

5 '-monophosphate (GMP), began with studies of nitrogen metabolism in birds. The
major end product of nitrogen metabolism in birds and some reptiles is uric acid
(Figure 18.1) rather than urea, as in mammals. Researchers in the 1950s discovered that
uric acid and nucleic acid purines arise from the same precursors and reaction sequence.
Homogenates of pigeon liver — a tissue in which purines are actively synthesized — were
a convenient source of enzymes for studying the steps in purine biosynthesis. The path-
way in avian liver has since been found in many other organisms.

When isotopically labeled compounds such 13 C0 2 , H 13 COCT(formate), and
+ H 3 N-CH 2 — 13 COO _ (glycine) were administered to pigeons and rats the result was ex-
cretion of labeled uric acid. This uric acid was isolated and chemically degraded to
identify the positions of the labeled carbon and nitrogen atoms. The carbon from car-
bon dioxide was incorporated into C-6, and the carbon from formate into C-2 and C-8
of purines. Ultimately, the sources of the ring atoms were shown to be N-l, aspartate;
C-2 and C-8, formate via 10-formyltetrahydrofolate (Section 7.9); N-3 and N-9, amide
groups from glutamine; C-4, C-5, and N-7, glycine; and C-6, carbon dioxide. These
findings are summarized in Figure 18.2.

The purine ring structure is not synthesized as a free base but as a substituent of ribose
5-phosphate. The ribose 5-phosphate for purine biosynthesis comes from 5-phosphoribosyl
1- pyrophosphate (PRPP) also known as 5-phosphoribosyl 1 -diphosphate. PRPP is syn-
thesized from ribose 5 -phosphate and ATP in a reaction catalyzed by ribose-phosphate
diphosphokinase (Figure 18.3); PRPP then donates ribose 5-phosphate to serve as the
foundation on which the purine structure is built. PRPP is also a precursor for the biosyn-
thesis of pyrimidine nucleotides, although in that pathway it reacts with a preformed
pyrimidine to form a nucleotide. PRPP is also used in the nucleotide salvage pathways
and in the biosynthesis of histidine (Figure 17.23).

The initial product of the purine nucleotide biosynthetic pathway is inosine 5'-
monophosphate (IMP, or inosinate) (Figure 18.4), a nucleotide containing hypoxan-
thine (6-oxopurine) as its base. The ten-step pathway for the de novo synthesis of IMP
was discovered in the 1950s by the research groups of John M. Buchanan and G. Robert
Greenberg. The painstaking isolation and structural characterization of the intermediates
took about ten years.

The pathway to IMP is shown in Figure 18.5. It begins with displacement of the py-
rophosphoryl group of PRPP by the amide nitrogen of glutamine in a reaction catalyzed
by glutamine-PRPP amidotransferase. Note that the configuration of the anomeric
carbon is inverted from a to /3 in this nucleophilic displacement — the /3 configuration
persists in completed purine nucleotides. The amino group of the product, phosphoribo-
sylamine, is then acylated by glycine to form glycinamide ribonucleotide. The mechanism
of this reaction, in which an enzyme-bound glycyl phosphate is formed, resembles that of
glutamine synthetase that has y- glutamyl phosphate as an intermediate (Reaction 10.17).

Step 3 consists of transfer of a formyl group from 10-formyltetrahydrofolate to the
amino group destined to become N-7 of IMP. In step 4, an amide is converted to an ami-
dine (RHN— C=NH) in an ATP- dependent reaction in which glutamine is the nitrogen
donor. Step 5 is a ring- closure reaction that requires ATP and produces an imidazole de-
rivative. C0 2 is incorporated in step 6 by attachment to the carbon that becomes C-5 of
IMP. This carboxylation is unusual because it does not require biotin. Bicarbonate is first
attached, in an ATP-dependent step, to the amino group that becomes N-3 of IMP. The
carboxylated intermediate then undergoes a rearrangement in which the carboxylate
group is transferred to the carbon atom that becomes C-5 of the purine ring (Figure 18.6).
These steps are catalyzed by two separate proteins in Escherichia coli but in eukaryotes
they are catalyzed by a multifunctional enzyme. Vertebrate versions of this enzyme trans-
fer the carboxylate group directly to the final position in carboxyaminoimidazole ribonu-
cleotide (CAIR). The vertebrate enzymes are much more efficient. There doesn’t seem to
be any reason why the enzymes in other species have to undergo a two-step reaction.

The amino group of aspartate is incorporated into the growing purine ring system
in steps 7 and 8. First, aspartate condenses with the newly added carboxylate group to
form an amide, specifically a succinylocarboxamide. Then adenylosuccinate lyase cat-
alyzes a nonhydrolytic cleavage reaction that releases fumarate. This two-step process

Uric acid

▲ Figure 18.1
Uric acid.

0 0

1 y

^ 1/3

0 O — P=0

▲ Structure of adenosine triphosphate (ATP).

The nitrogenous base adenine (blue) is at-
tached to ribose (black). Three phosphoryl
groups (red) are bound to the ribose at the
5' position.

Ribose 5-phosphate is produced by
the pentose phosphate pathway
(Section 12.4).

THE MAJOR PURINES

Adenine

(6-Aminopurine)

Guanine

(2-Amino-6-oxopurine)
▲ Adenine, Guanine

552 CHAPTER 18 Nucleotide Metabolism

Aspartate

C0 2 Glycine

nT 6 sC-

\ /

Glutamine

1 0-Formyltetrahydrofolate

▲ Figure 18.2

Sources of the ring atoms in purines synthesized
de novo.

Ribose 5-phosphate
(a anomer)

Ribose-phosphate

diphosphokinase

AMP^

AMP

O— P—O— P—O

▲ G. Robert Greenberg (1918-2005).

Greenberg’s research group worked out
many of the reactions of the purine biosyn-
thesis pathway.

▲ Figure 18.4

Inosine 5 -monophosphate (IMP, or inosinate).

IMP is converted to other purine nucleotides.
Much of the IMP is degraded to uric acid in
birds and primates.

5-Phospho-u-D-ribosyl 1 -pyrophosphate
(PRPP)

▲ Figure 18.3

Synthesis of 5-phosphoribosyl 1 -pyrophosphate (PRPP) from ribose 5-phosphate and ATP. Ribose-
phosphate diphosphokinase catalyzes the transfer of a pyrophosphoryl group from ATP to the
1-hydroxyl oxygen of ribose 5-phosphate.

results in the transfer of an amino group containing the nitrogen destined to become
N-l of IMP. The two steps are similar to steps 2 and 3 of the urea cycle (Figure 17.43)
except that in this case ATP is cleaved to ADP + Pj rather than to AMP + PPi.

In step 9, which resembles step 3, the cosubstrate 10-formyltetrahydro folate donates
a formyl group ( — CH = 0) to the nucleophilic amino group of aminoimidazole car-
boxamide ribonucleotide. The amide nitrogen of the final intermediate then condenses
with the formyl group in a ring closure that completes the purine ring system of IMP.

The synthesis of IMP consumes considerable energy. ATP is converted to AMP dur-
ing the synthesis of PRPP and steps 2, 4, 5, 6, and 7 are driven by the conversion of ATP
to ADR Additional ATP is required for the synthesis of glutamine from glutamate and
ammonia (Figure 17.4).

BOX 18.1

COMMON NAMES OF THE BASES

Adenine

from the Greek adenas , “gland”: first isolated from pancreatic
glands (1885)

Cytosine

derived from cyto- from the Greek word for “receptacle,” referring
to cells (1894)

Guanine

originally isolated from “guano” or bird excrement (1850)

Uracil

origin uncertain, possibly from “urea” (1890)

Thymine

first isolated from thymus glands (1894)

Xanthine

from the Greek word for “yellow” (1857)

©,

0,POCH,

O— P—0— P—0

,©

OH OH 0° 0 G
5-Phospho-a-D-ribosyl 1 -pyrophosphate (PRPP)

Glutamine —

Glutamate

H 2 0

Glutamine-PRPP

amidotransferase

► PP:

h 2 o

^ 2 Pj

5-Phospho-/3-D-ribosylamine (PRA)

^-n H3

-° e (D

Glycine

ATP

GAR synthetase
ADP + P:

H 2 cr

.nh 2

R5'P

Glycinamide ribonucleotide (GAR)

0 s

10-Formyl-
tetra hydrofolate

Tetrahydrofolate

GAR transformylase

H 2^y°

8 C.

R5'P

KEY CONCEPT

Nucleotide biosynthesis pathways are
energetically expensive.

COO*

i©

ribonucleotide

(SAICAR)

©O"

H,N'

*-^-N

II >CH
• /

"N
R5'P

Carboxyaminoimidazole
ribonucleotide (CAIR)

2H©<

HCOo

ADP + Pj

AIR carboxylase

ATP

hc r\

II BCH

3 Cl 9

R5'P

Formylglycinamide ribonucleotide (FGAR) Aminoimidazole ribonucleotide (AIR)

Glutamine

Glutamate

AIR synthetase

H 2 N 1

H,N'

II >CH

Cl 9 /

R5'P

Aminoimidazole carboxamide
ribonucleotide (AICAR)

10-Formyl-

tetrahydrofolate

Tetrahydrofolate

H 2 N i

AICAR

transformylase

-N

sCH

9 /

R5'P

Formamidoimidazole carboxamide
ribonucleotide (FAICAR)

HoO

HN i
HC

IMP

cyclohydrolase

sC-

-N

sCH

9 /

R5'P

Formylglycinamidine ribonucleotide (FGAM)

R5'P

Inosine 5'-monophosphate (IMP)

▲ Figure 18.5

Ten-step pathway for the de novo synthesis of IMP. R5'P stands for ribose 5'-phosphate. The atoms are numbered according to their positions in the
completed purine ring structure.

553

554 CHAPTER 18 Nucleotide Metabolism

Aminoimidazole

ribonucleotide

(AIR)

Vertebrates

A/-Carboxya mi no imidazole
ribonucleotide

▲ Figure 18.6

/V-Carboxyaminoimidazole ribonucleotide is sometimes an intermediate in the conversion of AIR to CAIR.

Carboxyaminoimidazole

ribonucleotide

(CAIR)

▲ John M. (“Jack”) Buchanan (1917-2007).

Buchanan’s group discovered many of the
purine biosynthesis pathway reactions. He
and Greenberg were friendly competitors
sharing many of their research results.

18.2 Other Purine Nucleotides Are Synthesized
from IMP

IMP can be converted to AMP or GMP (Figure 18.7). Two enzymatic reactions are re-
quired for each of these conversions. AMP and GMP can then be phosphorylated to
their di- and triphosphates by the actions of specific nucleotide kinases (adenylate ki-
nase and guanylate kinase, respectively) and the broadly specific nucleoside diphos-
phate kinase (Section 10.6).

The two steps that convert IMP to AMP closely resemble steps 7 and 8 in the
biosynthesis of IMP. First, the amino group of aspartate condenses with the keto group
of IMP in a reaction catalyzed by GTP- dependent adenylosuccinate synthetase. Next,
the elimination of fumarate from adenylosuccinate is catalyzed by adenylosuccinate
lyase, the same enzyme that catalyzes step 8 of the de novo pathway to IMP.

The first step in the conversion of IMP to GMP is the oxidation of C-2 catalyzed by
NAE® -dependent IMP dehydrogenase. This reaction proceeds by the addition of a
molecule of water to the double bond between C-2 and N-3 followed by oxidation of
the hydrate. The product of the oxidation is xanthosine monophosphate (XMP). Next,
the amide nitrogen of glutamine replaces the oxygen at C-2 of XMP in an ATP-dependent
reaction catalyzed by GMP synthetase. The use of GTP as a cosubstrate in the synthesis
of AMP from IMP, and of ATP in the synthesis of GMP from IMP, helps balance the for-
mation of the two products.

Purine nucleotide synthesis is regulated in cells by feedback inhibition. Several
enzymes that catalyze steps in the biosynthesis of purine nucleotides exhibit allosteric be-
havior in vitro. Ribose-phosphate diphosphokinase is inhibited by several purine ribonu-
cleotides but only at concentrations higher than those usually found in the cell. PRPP is a
donor of ribose 5 -phosphate in more than a dozen reactions so we would not expect
PRPP synthesis to be regulated exclusively by the concentrations of purine nucleotides.
The enzyme that catalyzes the first committed step in the pathway of purine nucleotide
synthesis, glutamine-PRPP amidotransferase (step 1 in Figure 18.5), is allosterically in-
hibited by 5 '-ribonucleotide end products (IMP, AMP, and GMP) at intracellular concen-
trations. This step appears to be the principal site of regulation of this pathway.

The paths leading from IMP to AMP and from IMP to GMP also appear to be reg-
ulated by feedback inhibition. Adenylosuccinate synthetase is inhibited in vitro by AMP,
the product of this two-step branch. Both XMP and GMP inhibit IMP dehydrogenase.
The pattern of feedback inhibition in the synthesis of AMP and GMP is shown in
Figure 18.8. Note that the end products inhibit two of the initial common steps as well
as steps leading from IMP at the branch point.

18.3 Synthesis of Pyrimidine Nucleotides 555

18.3 Synthesis of Pyrimidine Nucleotides

Uridine 5 '-monophosphate is the precursor of other pyrimidine nucleotides. The path-
way for the biosynthesis of UMP is simpler than the purine pathway and consumes
fewer ATP molecules. The pyrimidine ring atoms come from bicarbonate that contributes
C-2; the amide group of glutamine (N-3); and aspartate that contributes the remaining
atoms (Figure 18.9). C-2 and N-3 are incorporated after formation of the intermediate
carbamoyl phosphate.

PRPP is required for the biosynthesis of pyrimidine nucleotides but the sugar-
phosphate from PRPP is donated after the ring has formed rather than entering the

Aspartate

Adenylosuccinate

synthetase

GTP
GDP +

OH OH
IMP

NAD 0

r H2 °

NADH + H® ^

IMP dehydrogenase

|_|

°ooc— ch 2 — c— coo®

Adenylosuccinate

lyase

Fumarate

Xanthosine monophosphate
(XMP)

H 2 0 + ATP — v Glutam

GMP synthetase

PPj + AMP Glutamate

\ f

▲ Figure 18.7

Pathways for the conversion of IMP to AMP and to GMP.

556 CHAPTER 18 Nucleotide Metabolism

Figure 18.8 ►

Feedback inhibition in purine nucleotide
biosynthesis.

Ribose 5-phosphate

\ /

✓ i ■

i iii

Ribose-phosphate

i diphosphokinase

■|— ——————> «- — — — — — —

I ‘

r * [ - ^

| I PRPP

Glutamine-PRPP

amidotransferase

5-Phosphoribosylamine

(Steps 2-10)

Fumarate

▲ Adenylosuccinate lyase from E. coli. The

enzyme is a homodimer. One subunit is col-
ored blue and the other is purple. This is a
mutant enzyme (H171N) showing the two
products, AMP and fumarate, bound at the
active sites. Adenylosuccinate lyase cat-
alyzes similar steps in the IMP synthesis
pathway and in the conversion of IMP to
AMP. [PDB 2PTQ]

Glutamine

hco 3 °

Aspartate

▲ Figure 18.9

Sources of the ring atoms in pyrimidines.

The immediate precursor of C-2 and N-3
is carbamoyl phosphate.

v IMP

; Jk

Adenylosuccinate

synthetase

Adenylosuccinate

lyase

* \

dehydrogenase

XMP

GMP

synthetase

AMP

GMP

pathway in the first step. A compound with a completed pyrimidine ring — orotate
(6-carboxyuracil) — reacts with PRPP to form a pyrimidine ribonucleotide in the fifth
step of the six- step pathway.

A. The Pathway for Pyrimidine Synthesis

The six-step pathway for pyrimidine synthesis is shown in Figure 18.10. The first two
steps generate a noncyclic intermediate that contains all the atoms destined for the
pyrimidine ring. This intermediate, carbamoyl aspartate, is enzymatically cyclized. The
product is dihydroorotate and it is subsequently oxidized to orotate. Orotate is then
converted to the ribonucleotide orotidine 5 '-monophosphate (OMP, or orotidylate)
that undergoes decarboxylation to form UMP (uridylate). This pyrimidine nucleotide
is the precursor not only of all other pyrimidine ribonucleotides but also of the pyrimi-
dine deoxyribonucleotides. The enzymes required for pyrimidine synthesis are organ-
ized and regulated differently in prokaryotes and eukaryotes.

The first step in the pathway of pyrimidine biosynthesis is the formation of carbamoyl
phosphate from bicarbonate plus the amide nitrogen of glutamine and ATP. This reaction
is catalyzed by carbamoyl phosphate synthetase (or by carbamoyl phosphate synthetase II
activity in mammalian cells). It requires two molecules of ATP — one to drive formation of
the C — N bond and the other to donate a phosphoryl group. This enzyme is not the same
carbamoyl phosphate synthetase that is used in the urea cycle. That enzyme, carbamoyl
phophate synthetase I, assimilates free ammonia whereas this enzyme (carbamoyl phos-
phate synthetase II in animals) transfers an amino group from glutamine.

The activated carbamoyl group of carbamoyl phosphate is transferred to aspartate
to form carbamoyl aspartate in the second step of UMP biosynthesis. This reaction is
catalyzed by a famous enzyme, aspartate transcarbamoylase (ATCase). The mechanism
involves the nucleophilic attack of the aspartate nitrogen on the carbonyl group of
carbamoyl phosphate.

Dihydroorotase catalyzes the third step of UMP biosynthesis — the reversible closure
of the pyrimidine ring (Figure 18.10). The product, dihydroorotate, is then oxidized by the

18.3 Synthesis of Pyrimidine Nucleotides 557

coa

»©

HoN — C — H

ChH 2

H ? N

Glutamine

HCO

Carbamoyl
CD phosphate
synthetase

Glutamate

2 ATP + H,0

2ADP+ P

11 ©

H 2 N — c— OPOb^

Carbamoyl phosphate

'\ H

COO v

Carbamoyl aspartate
(3) Dihydroorotase

l-UO

A H

COO v

N O

Uridine 5'-monophosphate
(UMP)

HC0 3 °^

h 2 o -

OMP decarboxylase

HN'

CO o

Orotidine 5'-monophosphate
(OMP)

2 Pi ^

H-,0

PPi

PRPP

Orotate

phosphoribosyl- (5)
transferase

,©

L-Dihydroorotate

Dihydroorotate

dehydrogenase

QH 2

'N'

COO

Orotate

▲ Figure 18.10

Six-step pathway for the synthesis of UMP in prokaryotes. In eukaryotes, steps 1 through 3 are
catalyzed by a multifunctional protein called dihydroorotate synthase, and steps 5 and 6 are
catalyzed by a bifunctional enzyme, UMP synthase.

action of dihydroorotate dehydrogenase to form orotate. In eukaryotes, dihydroorotate is
produced in the cytosol by steps 1 through 3. It then passes through the outer mito-
chondrial membrane prior to being oxidized to orotate by the action of dihydroorotate
dehydrogenase. This enzyme is associated with the inner mitochondrial membrane. Its
substrate binding site is located on the outer surface. The enzyme is an iron-containing

THE MAJOR PYRIMIDINES
NH 2

Cytosine

(2-Oxo-4-aminopyrimidine)

Thymine

(2,4-Dioxo-5-methylpyrimidine)

Uracil

(2,4-Dioxopyrimidine)

▲ Cytosine, Thymine, Uracil

558

CHAPTER 18 Nucleotide Metabolism

Orotidine 5'-phosphate decarboxylase
(OMP decarboxylase) is one of the
most efficient enzymes known
(Table 5.2).

flavoprotein that catalyzes the transfer of electrons to ubiquinone (Q) forming ubiquinol
(QH 2 ). Electrons from QH 2 are then transferred to 0 2 via the electron transport chain.

Once formed, orotate displaces the pyrophosphate group of PRPP, producing OMP
in a reaction catalyzed by orotate phosphoribosyltransferase. The subsequent hydrolysis
of pyrophosphate makes this reaction essentially irreversible.

Finally, OMP is decarboxylated to form UMP in a reaction catalyzed by OMP de-
carboxylase. In eukaryotes, orotate produced in the mitochondria moves to the cytosol
where it is converted to UMP. A bifunctional enzyme known as UMP synthase catalyzes
both the reaction of orotate with PRPP to form OMP and the rapid decarboxylation of
OMP to UMP.

In mammals, the intermediates formed in steps 1 and 2 (carbamoyl phosphate and
carbamoyl aspartate) and OMP (from step 5) are not normally released to solvent but
remain bound to enzyme complexes and are channeled from one catalytic center to the
next. Several multifunctional proteins, each catalyzing several steps, also occur in the
pathway of purine nucleotide biosynthesis in some organisms.

BOX 18.2 HOW SOME ENZYMES TRANSFER AMMONIA FROM GLUTAMINE

Several enzymes that use glutamine as an amide donor have a
molecular tunnel running through the interior of the protein.
This is an example of metabolite channeling (Section 5.11).
Carbamoyl phosphate synthetase from E. coli is the most
fully studied of these enzymes. It catalyzes the synthesis of
carbamoyl phosphate from bicarbonate and glutamine:

Glutamine + HC0 3 ° + 2 ATP + H 2 Q

Carbamoyl
phosphate
synthetase II

from the glutamine-binding site, where a molecule of ammo-
nia is released from glutamine, to the second ATP-binding
site, where ammonia is carboxylated, and finally to the third
site where carbamoyl phosphate is formed. Ammonia that is
released from glutamine at the active site in the small subunit
does not equilibrate with solvent but proceeds down the tun-
nel and undergoes the reactions that eventually produce car-
bamoyl phosphate. Several of the intermediates in the overall
reaction are quite unstable and would be degraded by water
if they were not protected by being in a tunnel.

Carbamoyl phosphate formed in this reaction is
used in the synthesis of pyrimidine nucleotides.
(A different carbamoyl phosphate synthetase
that uses ammonia rather than glutamine as its
substrate is discussed in Section 17.7A.)

Carbamoyl phosphate synthetase of E. coli is
a heterodimer with one small subunit and one
large subunit (see figure). The synthesis of car-
bamoyl phosphate from glutamine proceeds via
three intermediates, each formed at a different
active site. ATP reacts at two of these sites. The
three sites are connected by a tunnel that runs

► Carbamoyl phosphate synthetase from E. coli. The

small subunit (/V-terminal domain, purple) contains
the active site for glutamine hydrolysis releasing
NH 3 . The large subunit is shown in blue. NH 3 is
converted to the unstable intermediate carbamate
(H 2 N — COOH) at its upper ATP-binding site. Carba-
mate is then phosphorylated at the C-terminal
(lower) ATP-binding site. A molecule of ADP is
bound in each ATP-binding site. The molecular
tunnel connecting the three active sites is shown
by the thick blue wire. [PDP 1A9X]

Gin-binding site

ATP-binding site

18.4 CTP Is Synthesized from UMP 559

B. Regulation of Pyrimidine Synthesis

Regulation of pyrimidine biosynthesis also differs between prokaryotes and eukaryotes.
Although the six enzymatic steps leading to UMP are the same in all species, the enzymes
are organized differently in different organisms. In E. coli , each of the six reactions is cat-
alyzed by a separate enzyme. In eukaryotes, a multifunctional protein in the cytosol
known as dihydroorotate synthase contains separate catalytic sites (carbamoyl phosphate
synthetase II, ATCase, and dihydroorotase) for the first three steps of the pathway.

In addition to being an intermediate in pyrimidine synthesis, carbamoyl phosphate
is a metabolite in the pathway for the biosynthesis of arginine via citrulline (Figure
17.43). The same carbamoyl phosphate synthetase in prokaryotes is also used in both
pyrimidine and arginine biosynthetic pathways. This enzyme is allosterically inhibited by
pyrimidine ribonucleotides such as UMP, the product of the pyrimidine biosynthetic path-
way. It is activated by L-ornithine, a precursor of citrulline, and by purine nucleotides,
the substrates (along with pyrimidine nucleotides) for the synthesis of nucleic acids.
Eukaryotic carbamoyl phosphate synthetase II is also allosterically regulated. PRPP and
IMP activate the enzyme and several pyrimidine nucleotides inhibit it.

The next enzyme of the pathway is aspartate transcarbamoylase (ATCase). ATCase
from E. coli is the most thoroughly studied allosteric enzyme. ATCase catalyzes the first
committed step of pyrimidine biosynthesis since carbamoyl phosphate can enter pathways
leading either to pyrimidines or to arginine in bacteria. This enzyme is inhibited by pyrim-
idine nucleotides and activated in vitro by ATP. ATCase in E. coli is only partially inhibited
(50% to 70%) by the most potent inhibitor, CTP, but inhibition can be almost total when
both CTP and UTP are present. UTP alone does not inhibit the enzyme. The allosteric
controls — inhibition by pyrimidine nucleotides and activation by the purine nucleotide
ATP — provide a means for carbamoyl phosphate synthetase and ATCase to balance the
pyrimidine nucleotide and purine nucleotide pools in E. coli. The ratio of the concentra-
tions of the two types of allosteric modulators determines the activity level of ATCase.

E. coli ATCase has a complex structure with binding sites for substrates and allosteric
modulators on separate subunits. The enzyme contains six catalytic subunits arranged as two
trimers and six regulatory subunits arranged as three dimers (Figure 18. 1 1 ). Each subunit of a
catalytic trimer is connected to a subunit of the other catalytic trimer through a regulatory
dimer. When one molecule of aspartate binds, in the presence of carbamoyl phosphate, all six
catalytic subunits change to a conformation having increased catalytic activity.

Eukaryotic ATCase is not feedback- inhibited. Regulation by feedback inhibition is not
necessary because the pyrimidine pathway can be controlled by regulating the enzyme pre-
ceding ATCase, carbamoyl phosphate synthetase II. The substrate of ATCase in eukaryotes
is not a branch-point metabolite — the synthesis of carbamoyl phosphate and citrulline for
the urea cycle occurs in mitochondria, and the synthesis of carbamoyl phosphate for
pyrimidines occurs in the cytosol. The pools of carbamoyl phosphate are separate.

18.4 CTP Is Synthesized from UMP

UMP is converted to CTP in three steps. Uridylate kinase (UMP kinase) catalyzes the
transfer of the y-phosphoryl group of ATP to UMP to generate UDP, and then nucleo-
side diphosphate kinase catalyzes the transfer of the y-phosphoryl group of a second
ATP molecule to UDP to form UTP. Two molecules of ATP are converted to two mole-
cules of ADP during the synthesis of UTP from UMP.

ATP ADP ATP ADP

ump a — UDP

CTP synthetase then catalyzes the ATP-dependent transfer of the amide nitrogen from
glutamine to C-4 of UTP forming CTP (Figure 18.12). This reaction is chemically anal-
ogous to step 4 of purine biosynthesis (Figure 18.5) and to GMP synthesis from XMP
catalyzed by GMP synthetase (Figure 18.7).

CTP synthetase is allosterically inhibited by its product, CTP, and in E. coli it is al-
losterically activated by GTP (Figure 18.13). The regulation of ATCase and CTP syn-
thetase balances the concentrations of endogenous pyrimidine nucleotides. Elevated
levels of CTP block further synthesis of CTP by inhibiting CTP synthetase. Under these

Active

site

CTP

▲ Figure 18.11

ATCase from Escherichia coli. The top struc-
ture has two regulatory subunits (purple)
with a bound CTP. The two catalytic sub-
units (blue) have a bound substrate analog
that identifies the active site. Note the large
distance between the allosteric site where
CTP binds and the active site of the en-
zyme. Three of these units are bound to-
gether to produce a large hexameric ring
(below) and two of these hexameric rings
stack together to create the complete
12-subunit enzyme. [PDB 2FZC (top) 9ATC
(bottom)].

560 CHAPTER 18 Nucleotide Metabolism

H 2 0 + ATP

CTP synthetase

P; + ADP

Glutamine

Glutamate

▲ Figure 18.12
Conversion of UTP to CTP.

Aspartate ATp

+ i

Carbamoyl phosphate

i +

N/ ATCase

Carbamoyl aspartate

De novo pathway
(Steps 3-6)

UMP

UDP

UTP

CTP synthetase

▲ Figure 18.13

Regulation of pyrimidine nucleotide synthesis
in E. coli. Allosteric regulation of ATCase and
CTP synthetase by both purine and pyrimi-
dine nucleotides helps balance nucleotide
synthesis.

conditions, UMP synthesis will be slowed but not stopped since CTP only partially
inhibits ATCase. UMP can still be used in RNA synthesis and as a precursor to dTTP
(Section 18.6). ATCase is completely inhibited when the concentrations of both UTP
and CTP are elevated. Elevated concentrations of the purine nucleotides ATP and GTP
increase the rates of synthesis of the pyrimidine nucleotides and this helps balance the
supplies of purine and pyrimidine nucleotides.

18.5 Reduction of Ribonucleotides
to Deoxyribonucleotides

The 2'-deoxyribonucleoside triphosphates are synthesized by the enzymatic reduction of
ribonucleotides. This reduction occurs at the nucleoside diphosphate level in most organ-
isms. Peter Reichard and his colleagues showed that all four ribonucleoside diphosphates —
ADP, GDP, CDP, and UDP — are substrates of a single, closely regulated, ribonucleoside
diphosphate reductase. In some microorganisms, including species of Lactobacillus,
Clostridium , and Rhizobium , ribonucleoside triphosphates are the substrates for reduction
by a cobalamin-dependent reductase. Both types of enzymes are called ribonucleotide re-
ductase (class I and class II, respectively), although the more precise names are ribonucleo-
side diphosphate reductase and ribonucleoside triphosphate reductase.

NADPH provides the reducing power for the synthesis of deoxyribonucleoside
diphosphates in class I enzymes. A disulfide bond at the active site of ribonucleotide reduc-
tase is reduced to two thiol groups that reduce C-2' of the ribose moiety of the nucleotide
substrate by a complex free-radical mechanism. As shown in Figure 18.14, electrons are
transferred from NADPH to ribonucleotide reductase via the flavoprotein thioredoxin re-
ductase and the dithiol protein coenzyme thioredoxin (Figure 7.35). Thioredoxin reductase
of prokaryotes and yeast has a dithiol/disulfide (cysteine pair) group in the active site. In
mammalian thioredoxin reductase, the oxidation-reduction center differs by having one
residue of cysteine and one of selenocysteine. Once formed, dADP, dGDP, and dCDP are
phosphorylated to the triphosphate level by the action of nucleoside diphosphate kinases.
dUDP, as we will see in the next section, is converted to dTMP via dUMP. A third version of
ribonucleotide reductase (class III) uses 5-adenosylmethionine as a cofactor.

Ribonucleotide reductase has a complicated mechanism of allosteric regulation
that supplies a balanced pool of the deoxynucleotides required for DNA synthesis. Both
the substrate specificity and the catalytic rate of ribonucleotide reductase are regulated
in eukaryotic cells by the reversible binding of nucleotide metabolites. The allosteric
modulators — ATP, dATP, dTTP, and dGTP — act by binding to ribonucleotide reductase
at either of two regulatory sites. One allosteric site, called the activity site , controls the
activity of the catalytic site. A second allosteric site, called the specificity site , controls the
substrate specificity of the catalytic site (Figure 18.15). The binding of ATP to the activity
site forms an activated enzyme whereas the binding of dATP to the activity site inhibits
all enzymatic activity. When ATP is bound to the activity site and either ATP or dATP is
bound to the specificity site, the reductase becomes pyrimidine specific, catalyzing the
reduction of CDP and UDP. The binding of dTTP to the specificity site activates the re-
duction of GDP, and the binding of dGTP activates the reduction of ADP. The allosteric
regulation of ribonucleotide reductase, summarized in Table 18.1, controls enzyme ac-
tivity and ensures a balanced selection of deoxyribonucleotides for DNA synthesis.

18.6 Methylation of dUMP Produces dTMP

Deoxythymidylate (dTMP) is formed from UMP in four steps. UMP is phosphorylated
to UDP that is reduced to dUDP and dUDP is dephosphorylated to dUMP. dUMP is
then methylated to dTMP.

UMP > UDP > dUDP > dUMP > dTMP (18.2)

The conversion of dUDP to dUMP can occur by two routes. dUDP can react with
ADP in the presence of a nucleoside monophosphate kinase to form dUMP and ATP.

dUDP + ADP dUMP + ATP

(18.3)

18.6 Methylation of dUMP Produces dTMP 561

Ribonucleoside

diphosphate

Thioredoxin reductase

Deoxyribonucleoside

diphosphate

▲ Figure 18.14

Reduction of ribonucleoside diphosphates. Three proteins are involved: the NADPH-dependent flavoprotein thioredoxin reductase, thioredoxin, and
ribonucleotide reductase. B represents a purine or pyrimidine base. S(e) represents either sulfur or selenium.

dUDP can also be phosphorylated to dUTP at the expense of ATP through the action of
nucleoside diphosphate kinases. dUTP is then rapidly hydrolyzed to dUMP + PPj by the
action of deoxyuridine triphosphate diphosphohydrolase (dUTPase).

dUDP + ATP

* dUTP

ADP

H,0

dUMP + PPi

(18.4)

The rapid hydrolysis of dUTP prevents it from being accidentally incorporated into
DNAin place of dTTP.

dCMP can also be a source of dUMP via hydrolysis catalyzed by dCMP deaminase.

dCMP + H 2 0 * dUMP + HN 4 @ (18.5)

The conversion of dUMP to dTMP is catalyzed by the enzyme known as thymidylate
synthase. (Because thymine occurs almost exclusively in DNA, the trivial names thymi-
dine and thymidylate are commonly used instead of deoxythymidine and deoxythymidy-
late.) 5,10-Methylenetetrahydrofolate is the donor of the one-carbon group in this reaction
(Figure 18.16). The carbon-bound methyl group (C — CH 3 ) in dTMP is more reduced
than the nitrogen-bridged methylene group (N — CH 2 — N) in 5,10-methylenetetrahy-
drofolate, whose oxidation state is equivalent to that of a nitrogen-bound hydroxymethyl
group (N — CH 2 OH) or formaldehyde. Thus, not only is methylenetetrahydrofolate a
coenzyme donating a one-carbon unit but it is also the reducing agent for the reaction,
furnishing a hydride ion and being oxidized to 7, 8 -dihydrofolate in the process. This is the
only known reaction in which the transfer of a one-carbon unit from a tetrahydrofolate
derivative results in its oxidation at N-5 and C-6 to produce dihydrofolate.

Table 18.1 Allosteric regulation of eukaryotic ribonucleotide reductase

Ligand bound
to activity site

Ligand bound
to specificity site

Activity of
catalytic site

dATP

Enzyme inactive

ATP

ATP or dATP

Specific for CDP or UDP

ATP

dTTP

Specific for GDP

ATP

dGTP

Specific for ADP

The structure of selenocysteine,
the 22nd amino acid, is shown in
Section 3.3.

▲ Peter Reichard (1925-).

Reichard worked for many years at the
Karolinska Institute in Sweden. In addition
to working on ribonucleotide reductase, he
was an active member of the Nobel Commit-
tee that selects candidates to receive the
Nobel Prize.

562 CHAPTER 18 Nucleotide Metabolism

Figure 18.15 ►

Ribonucleotide reductase. The complete
enzyme is an a 2 fi 2 tetramer. The structure
shown here (from E. coli) shows only the a 2
dimer of catalytic subunits. The activity site
is occupied by an ATP analog. A molecule of
TTP is bound to the specificity site and a
molecule of GDP is bound at the active site.
[PDB 3R1R + 4R1R]

Activity site
Catalytic site

Specificity site

Catalytic site
Activity site

BOX 18.3 FREE RADICALS IN THE REDUCTION OF RIBONUCLEOTIDES

The ribonucleotide reductase reaction is an unusual reaction
because it proceeds by a free radical mechanism. The first
clue to the free radical nature of the reaction was the obser-
vation that the reductase from E. coli could be isolated with a
tyrosine residue in the free radical form. This was the first
free radical protein to be discovered. The role of the tyrosine
radical is to convert the thiol group of an active-site cysteine
residue to a thiyl radical. (In the Lactobacillus enzyme, cobal-
amin serves to convert the active-site thiol to a radical.)

The proposed mechanism is shown in the accompanying
figure. The active site of the reductase has three cysteine
residues — one forms the free radical and the other two are an
oxidation-reduction group. The thiyl radical removes a hy-
drogen atom from the C-3' position of the ribonucleotide
forming a substrate radical. This substrate radical is first de-
hydrated (losing the C-2' — OH) and then reduced by the
cysteine reduction pair. A hydrogen atom is returned to C-3',
regenerating the thiyl radical.

-h 2 o, -h©

18.6 Methylation of dUMP Produces dTMP 563

5, 1 0-Methylenetetrahydrofolate

7,8-Dihydrofolate

Di hydrofolate
reductase

Tetrahydrofolate

◄ Figure 18.16

Cycle of reactions in the synthesis of thymidy-
late (dTMP) from dUMP. Thymidylate synthase
catalyzes the first reaction of this cycle pro-
ducing dTMP. The other product of the reac-
tion, dihydrofolate, must be reduced by
NADPH in a reaction catalyzed by dihydrofo-
late reductase before a methylene group can
be added to regenerate 5,10-methylenete-
trahydrofolate. Methylenetetrahydrofolate is
regenerated in a reaction catalyzed by serine
hydroxymethyltransferase.

Dihydrofolate must be converted to tetrahydrofolate before the coenzyme can accept
another one-carbon unit for further transfer reactions. The 5,6 double bond of dihydrofo-
late is reduced by NADPH in a reaction catalyzed by dihydrofolate reductase. Serine hy-
droxymethyltransferase (Figure 17.16) then catalyzes the transfer of the /3-CH 2 OH group
of serine to tetrahydrofolate to regenerate 5, 10-methylenetetrahydro folate.

Thymidylate synthase and dihydrofolate reductase are distinct polypeptides in most
organisms but in protozoa the two enzyme activities are contained on the same polypep-
tide chain. The dihydrofolate product of the first reaction is channeled from the thymidylate
synthase active site to the dihydrofolate reductase active site. Charge-charge interactions
between a positively charged region on the surface of the bifunctional enzyme and the
negatively charged dihydro folate (recall that it contains several y - glutamate residues;
Section 7.11) steer the product to the next active site.

dTMP can also be synthesized via the salvage of thymidine (deoxythymidine), cat-
alyzed by ATP-dependent thymidine kinase.

ATP ADP

^ — A, dTMP

Deoxythymidine

(Thymidine)

Thymidine

kinase

(18.6)

564 CHAPTER 18 Nucleotide Metabolism

BOX 18.4 CANCER DRUGS INHIBIT dTTP SYNTHESIS

Since dTMP is an essential precursor of DNA, any agent that
lowers dTMP levels drastically affects cell division. Thymidy-
late synthase and dihydro folate reductase have been major
targets for anticancer drugs because rapidly dividing cells are
particularly dependent on the activities of these enzymes.

The inhibition of either or both of these enzymes blocks the
synthesis of dTMP and therefore the synthesis of DNA.

5-Fluorouracil, methotrexate, and Tomudex are effective
in combating some types of cancer. 5-Fluorouracil is con-
verted to its deoxyribonucleotide, 5-fluorodeoxyuridylate,

which binds tightly to thymidylate synthase inhibiting the
enzyme and bringing the three-reaction cycle shown in
Figure 18.16 to a halt. Methotrexate, an analog of folate, is a
potent, relatively specific inhibitor of dihydrofolate reductase
that catalyzes step 2 of the cycle shown in Figure 18.16. The
resulting decrease in tetrahydrofolate levels greatly dimin-
ishes the formation of dTMP since dTMP synthesis depends
on adequate concentrations of methylenetetrahydrofolate.
Tomudex is a folate-based inhibitor of human thymidylate
synthase that has been approved for the treatment of cancer.

COO® reductase with the substrate analog

methotrexate (red) and the cosubstrate

▲ 5-Fluorouracil, methotrexate, and Tomudex are drugs designed to inhibit thymidylate synthase NADPH (gold) bound in the active site,
and block the growth of rapidly dividing cells. [PDB 1DLS]

Radioactive thymidine is often used as a highly specific tracer for monitoring intracellular
synthesis of DNA because it enters cells easily and its principal metabolic fate is conversion
to thymidylate and incorporation into DNA.

▲ Salvage pathways are a form of biochemical
recycling.

18.7 Modified Nucleotides

DNA and RNA contain a number of modified nucleotides. The ones present in transfer
RNA are well known (Section 21.8B) but the modified nucleotides in DNA are just as im-
portant. Some of the more common modified bases in DNA are shown in Figure 18.17.
Most of them are only found in a few species or in bacteriophage while others are more
widespread.

We will encounter N 6 -methyladenine in the next chapter when we discuss restric-
tion endonucleases. 5-Methylcytosine is a common modified base in mammalian DNA
because it plays a role in chromatin assembly and the regulation of transcription. About
3% of all deoxycytidylate residues in mammalian DNA are modified to 5-methylcytidine.
The methylation occurs after DNA is synthesized and the modified residues are at CG
sequences. All of these modified nucleotides are made in situ by enzymes that act on one
of the four common nucleotides in the DNA molecule.

18.8 Salvage of Purines and Pyrimidines

Nucleic acids are degraded to mononucleotides, nucleosides, and eventually, heterocyclic
bases during normal cell metabolism (Figure 18.18). The catabolic reactions are catalyzed by

18.9 Purine Catabolism 565

Methylcytosine

ribonucleases, deoxyribonucleases, and a variety
of nucleotidases, nonspecific phosphatases, and
nucleosidases or nucleoside phosphorylases.

Some of the purine and pyrimidine bases
formed in this way are further degraded (e.g.,
purines are converted to uric acid and other ex-
cretory products) but a considerable fraction is
normally salvaged by direct conversion back to
5 '-mononucleotides. PRPP is the donor of the
5-phosphoribosyl moiety for salvage reactions.

The degradation pathways are part of fuel me-
tabolism in animals. Purines and pyrimidines
formed during digestion are more likely to be
degraded while those formed inside the cell are
usually salvaged. The recycling of intact bases
conserves cellular energy.

The degradation of purine nucleotides to their respective purines and their salvage
through reaction with PRPP are outlined in Figure 18.19. Adenine phosphoribosyl -
transferase catalyzes the reaction of adenine with PRPP to form AMP and PPj. The
hydrolysis of PP*, catalyzed by pyrophosphatase, renders the reaction metabolically irre-
versible. Hypoxanthine-guanine phosphoribosyltransferase catalyzes similar reactions —
the conversion of hypoxanthine to IMP and of guanine to GMP with formation of PPj.

Pyrimidines are salvaged by the action of orotate phosphoribosyltransferase, which
catalyzes step 5 of the biosynthesis pathway (Figure 18.10). This enzyme can also cat-
alyze the conversion of pyrimidines other than orotate to the corresponding pyrimidine
nucleotides.

Nucleotides and their constituents are interconverted by many reactions, some of
which we have seen already. The actions of phosphatases, nucleotidases, and nucleosi-
dases or nucleoside phosphorylases can release bases from nucleotides. Reactions cat-
alyzed by phosphoribosyltransferases or nucleoside phosphorylases can salvage the
bases and nucleosides by converting them to the nucleotide level. Bases that are not
salvaged can be catabolized. The interconversions of purine nucleotides and their con-
stituents are summarized in Figure 18.20, and the interconversions of pyrimidine
nucleotides and their constituents are summarized in Figure 18.21.

18.9 Purine Catabolism

ch 2 oh

ChhOH

5-Methylcytosine 5-Hydroxymethylcytosine 5-Hydroxymethyluracil

N 6 -Methyladenine

2-Aminoadenine

▲ Figure 18.17
Modified bases in DNA.

Nucleic acids

Nucleases

Mononucleotides

Nucleotidases

and

phosphatases

Nucleosides

Nucleosidases
or

nucleoside phosphorylases

Bases

Salvage
reactions ,

Catabolism

5'-Mono-

nucleotides

Degradation

products

Most free purine and pyrimidine molecules are salvaged but some are catabolized.
Birds, some reptiles, and primates (including humans) convert purine nucleotides to
uric acid or urate, which is then excreted. In birds and reptiles, amino acid catabolism
also leads to uric acid; in mammals, surplus nitrogen from amino acid catabolism is dis-
posed of in the form of urea. Birds and reptiles cannot further catabolize uric acid
(urate) but many organisms degrade urate to other products.

As shown in Figure 18.20, AMP can be broken down to hypoxanthine and GMP is
broken down to guanine. The hydrolytic removal of phosphate from AMP and GMP
produces adenosine and guanosine, respectively. Adenosine can be deaminated to ino-
sine by the action of adenosine deaminase. Alternatively, AMP can be deaminated to

▲ Figure 18.18
Breakdown of nucleic acids.

Adenine

phosphoribosyl

transferase

PRPP

AMP

Adenosine

Adenine

IMP

Inosine

Hypoxanthine-
guanine
phosphoribosyl
transferase

PRPP

Hypoxanthine

GMP

Guanosine

Guanine

◄ Figure 18.19

Degradation and salvage of purines.

566

CHAPTER 18 Nucleotide Metabolism

Amino-group

Reduction transfer

Amino-group

Oxidation transfer Reduction

OH H OH OH OH OH OH OH OH OH OH H

Triphosphate

Diphosphate

Monophosphate

Nucleoside

Base

dATP

ATP

GTP

dADP 4 -

ADP

GDP

dAMP

AMP

IMP

-> XMP

-> GMP

PRPP

Adenine Adenine Hypoxanthine

dGTP

■» dGDP

dGMP

dGuanosine

\ V

Guanine

▲ Figure 18.20

Interconversions of purine nucleotides and their constituents. IMP, the first nucleotide product of the de novo biosynthetic pathway, is readily converted
to AMP and GMP, their di- and triphosphates, and the deoxy counterparts of these nucleotides. 5'-Phosphate groups are not shown in the abbrevi-
ated structures. [Adapted from Traut, T. W. (1988). Enzymes of nucleotide metabolism: the significance of subunit size and polymer size for biologi-
cal function and regulatory properties. Crit. Rev. Biochem. 23:121-169.]

IMP by the action of AMP deaminase and then IMP can be hydrolyzed to inosine. The
phosphorolysis of inosine produces hypoxanthine and the phosphorolysis of guanosine
produces guanine. Both these reactions (as well as the phosphorolysis of several de-
oxynucleosides) are catalyzed by purine-nucleoside phosphorylase and produce u-D-ribose
1 -phosphate (or deoxyribose 1 -phosphate) and the free purine base.

(Deoxy) Nucleoside + Pj Base + (Deoxy)-a-D-Ribose 1 -phosphate (18.7)

Adenosine is not a substrate of mammalian purine-nucleoside phosphorylase.

Hypoxanthine formed from inosine is oxidized to xanthine, and xanthine is oxidized
to urate (Figure 18.22). Either xanthine oxidase or xanthine dehydrogenase can catalyze
both reactions. Electrons are transferred to 0 2 to form hydrogen peroxide (H 2 0 2 ) in the
See Section 6.5D for a description of reactions catalyzed by xanthine oxidase. (The H 2 0 2 is converted to H 2 0 and 0 2 by the

the adenosine deaminase mechanism. action of catalase.) Xanthine oxidase is an extracellular enzyme in mammals and it appears

to be an altered form of the intracellular enzyme xanthine dehydrogenase that generates
the same products as xanthine oxidase but transfers electrons to NAD® to form NADH.
These two enzyme activities occur widely in nature and exhibit broad substrate specificity.
Their active sites contain complex electron- transfer systems that include an iron-sulfur
cluster, a pterin coenzyme with bound molybdenum, and FAD.

In most cells, guanine is deaminated to xanthine in a reaction catalyzed by guanase
(Figure 18.22). Animals that lack guanase excrete guanine. For example, pigs excrete
guanine but metabolize adenine derivatives further to allantoin, the major end product
of the catabolism of purines in most mammals.

Urate can be further oxidized in most organisms. Up until recently it was thought
that urate oxidase converted urate directly to allantoin but it is now known that the path-
way is more complex. The conversion of urate to the stereospecific product (S) -allantoin

18.9 Purine Catabolism 567

Reduction

X 11

Amino-group

transfer Reduction

nh 2 o

Methylation

1 1
cr^N

Triphosphate

Diphosphate

dCTP

dCDP <-

CTP <-

UTP

dUTP

Monophosphate

dUMP X

Nucleoside dCytidine

CDP

CMP

Cytidine

dTTP

dTDP

> dTMP

-> Uridine

dUridine dThymidine

Base

Cytosine

-> Uracil

Uracil

Thymine

▲ Figure 18.21

Interconversions of pyrimidine nucleotides and their constituents. UMP formed by the de novo pathway can be converted to cytidine and thymidine
phosphates, as well as to other uridine derivatives. 5'-Phosphate groups are not shown in the abbreviated structures. [Adapted from Traut, T. W.
(1988). Enzymes of nucleotide metabolism: the significance of subunit size and polymer size for biological function and regulatory properties.
Crit. Rev. Biochem. 23:121-169.]

requires urate oxidase plus two additional enzymes as shown in Figure 18.23. Peroxide
(H 2 0 2 ) and C0 2 are released in this series of reactions. Allantoin is the major end prod-
uct of purine degradation in most mammals (though not in humans, for whom the end
product is urate). It is also excreted by turtles, some insects, and also gastropods.

The enzyme allantoinase catalyzes hydrolytic opening of the imidazole ring of al-
lantoin to produce allantoate, the conjugate base of allantoic acid. Some bony fishes
(teleosts) possess allantoinase activity and excrete allantoate as the end product of
purine degradation.

H 2 0 + 0 2

H?0?

Xanthine

oxidase

f Xanthine
dehydrogenase I

Hypoxanthine u O + NAD 0 NADH + H 0

NH,

H 2 0

^Guanasej

Xanthine

H 2 0 + NAD©-

Xanthine dehydrogenase

NADH + H 0 ^

h 2 o + 0 2

Xanthine oxidase

^h 2 0 2

◄ Figure 18.22

Breakdown of hypoxanthine and guanine
to uric acid.

568 CHAPTER 18 Nucleotide Metabolism

°ooc

h 2 n

° H H

| \ = 0°

HUI

hydrolase

h 2 o

2-Oxo-4-hyd roxy-4-ca rboxy-
5-ureidoimidazoline (OHCU)

HINT

' N

-o

5-Hydroxyisourate

(HIU)

Urate oxidase

h 2 o 2 h 2 o o
+

0 2

C ^ \ Birds;

C = O some reptiles;
^ C ^ / prima tes

Uric acid

OHCU

QQ 2 decarboxylase

H 2 N

N^H

Most mammals; turtles;
some insects; gastropods

Allantoinase

(S)-Allantoin
- H,0

H ? N

coo v

x H I ^

C — N^-C-^-N — C

</ H H V

Allantoate

Some bony fishes

i©

cocr

O^H

Glyoxylate

H 2 0

2 H 2 0

Allantoicase

* 2 H 2 N — C — NH 2
Urea

Urease

2 H 2 0

Most fishes; amphibians;
freshwater mollusks

2 C0 2 + 4 NH 3

Plants; crustaceans; many
marine invertebrates

▲ Figure 18.23

Catabolism of uric acid through oxidation and hydrolysis. To the right of each compound are listed the organisms for which it is an excretory product.

▲ When they were alive, these snails could
convert urate to allantoin. Humans can’t do
that.

Most fishes, amphibians, and freshwater mollusks can further degrade allantoate.
These species contain allantoicase that catalyzes the hydrolysis of allantoate to one mol-
ecule of glyoxylate and two molecules of urea. Urea is the nitrogenous end product of
purine catabolism in these organisms.

Finally, several organisms — including plants, crustaceans, and many marine
invertebrates — can hydrolyze urea in a reaction catalyzed by urease. Carbon dioxide
and ammonia are the products of this reaction. Urease is found only in the cells of
organisms in which the hydrolysis of urea does not lead to ammonia toxicity. For exam-
ple, in plants, ammonia generated from urea is rapidly assimilated by the action of glut-
amine synthetase. In marine animals, ammonia is produced in surface organs such as
gills and is flushed away before it can accumulate to toxic levels. Most terrestrial organ-
isms would be poisoned by the final nitrogen-containing product, ammonia. The en-
zymes that catalyze urate catabolism have been lost through evolution by organisms
that excrete urate.

18.10 Pyrimidine Catabolism

The catabolism of pyrimidine nucleotides begins with hydrolysis to the corresponding nu-
cleosides and Pi, catalyzed by 5 '-nucleotidase (Figure 18.24). Initial hydrolysis to cytidine
can be followed by deamination to uridine in a reaction catalyzed by cytidine deaminase.

18.10 Pyrimidine Catabolism 569

BOX 18.5 LESCH-NYHAN SYNDROME AND GOUT

Defects in purine metabolism can have devastating effects. In
1964 Michael Lesch and William Nyhan described a severe
metabolic disease characterized by slow mental development,
palsylike spasticity, and a bizarre tendency toward self-
mutilation. Individuals afflicted with this disease, called
Lesch-Nyhan syndrome, rarely survive past childhood.
Prominent biochemical features of the disease are the excre-
tion of up to six times the normal amount of uric acid and a
greatly increased rate of purine biosynthesis.

The disease is caused by a hereditary deficiency of the
activity of the enzyme hypoxanthine-guanine phosphoribo-
syltransferase (Section 18.8). The deficiency is usually seen in
males because the mutation is recessive and the gene for this
enzyme is on the X chromosome. Lesch-Nyhan patients usu-
ally have less than 1% of the normal activity of the enzyme
and most show a complete absence of activity. In the absence
of hypoxanthine-guanine phosphoribosyltransferase, hypox-
anthine and guanine are degraded to uric acid instead of
being converted to IMP and GMP, respectively. The PRPP
normally used for the salvage of hypoxanthine and guanine
contributes to the synthesis of excessive amounts of IMP and
the surplus IMP is degraded to uric acid. It is not known how
this single enzyme defect causes the various behavioral
symptoms. The catastrophic effects of the deficiency indicate
that in some cells the purine salvage pathway in humans is
not just an energy-saving addendum to the central pathways
of purine nucleotide metabolism.

Gout is a disease caused by the overproduction or inade-
quate excretion of uric acid. Sodium urate is relatively insol-
uble and when its concentration in blood is elevated, it can
crystallize (sometimes along with uric acid) in soft tissues,
especially the kidney, and in toes and joints. Gout has several
causes including a deficiency of hypoxanthine-guanine
phosphoribosyltransferase activity resulting in less salvage of
purines and more catabolic production of uric acid. The dif-
ference between gout and Lesch-Nyhan syndrome is due to
the fact that gout patients retain up to 10% enzyme activity.
Gout can also be caused by defective regulation of purine
biosynthesis.

Sodium urate

Gout can be treated by giving patients allopurinol, a syn-
thetic C-7, N-8 positional isomer of hypoxanthine. Allopuri-
nol is converted in cells to oxypurinol, a powerful inhibitor
of xanthine oxidase. Administration of allopurinol prevents
the formation of abnormally high levels of uric acid. Hypox-
anthine and xanthine are more soluble than sodium urate
and uric acid and they are excreted when not reused by sal-
vage reactions.

Hypoxanthine

Xanthine

dehydrogenase

▲ Allopurinol and oxypurinol. Xanthine dehydrogenase catalyzes the oxidation of allopurinol,
an isomer of hypoxanthine. The product, oxypurinol, binds tightly to xanthine dehydrogenase,
inhibiting the enzyme.

The glycosidic bonds of uridine and thymidine are then cleaved by phosphorolysis in reactions
catalyzed by uridine phosphorylase and thymidine phosphorylase, respectively.
Deoxyuridine can also undergo phosphorolysis catalyzed by uridine phosphorylase. The
products of these phosphorolysis reactions are a-D-ribose 1 -phosphate or deoxyribose
1 -phosphate, thymine, and uracil.

The catabolism of pyrimidines ends with intermediates of central metabolism, so no
distinctive excretory products are formed. The breakdown of both uracil and thymine
involves several steps (Figure 18.24). First, the pyrimidine ring is reduced to a 5,6-dihy-
dropyrimidine in a reaction catalyzed by dihydrouracil dehydrogenase. The reduced ring

570 CHAPTER 18 Nucleotide Metabolism

Figure 18.24 ►

Catabolism of uracil and thymine.

him; 4

cr n

1 H

3CH

/<=^/ ch 3

Uracil

CT N

Dihydrouracil dehydrogenase

hn; 4 ;<r

J.CH
H

Thymine

NADPH + H e
^NADP @

CH-,

„CH,

HN^

.CHo

' N '

Dihydrouracil

H.O-

Dihydropyrimidinase

Dihydrothymine
-H 2 0

O CH,

H 2 N — C — N — CH 2 — CH 2 — COO
H

Ureidopropionate

h 2 o

h 3 n — ch 2 — ch 2 — coo'

/3-Alanine

Ureidopropionase

H 2 N — C — N — CH 2 — C — COO
H H

Ureidoisobutyrate
- H 2 0

CH,

,©

h 3 n — ch 2 — c— coo

/3-Aminoisobutyrate

,©

Acetyl CoA

Succinyl CoA

is then opened by hydrolytic cleavage of the N-3 — C-4 bond in a reaction catalyzed by
dihydropyrimidinase. The resulting carbamoyl-/3-amino acid derivative (ureidopropi-
onate or ureidoisobutyrate) is further hydrolyzed to NH 4 ®, HC0 3 ®, and a /3-amino
acid. /3-Alanine (from uracil) and /3-aminoisobutyrate (from thymine) can then be con-
verted to acetyl CoA and succinyl CoA, respectively, which can enter the citric acid cycle
and be converted to other compounds. In bacteria, /3-alanine can also be used in the syn-
thesis of pantothenate, a constituent of coenzyme A.

Problems 571

Summary

1. The synthesis of purine nucleotides is a ten-step pathway that
leads to IMP (inosinate). The purine is assembled on a founda-
tion of ribose 5-phosphate donated by 5-phosphoribosyl 1 -pyrophos-
phate (PRPP).

2. IMP can be converted to AMP or GMP.

3. In the six- step synthesis of the pyrimidine nucleotide UMP, PRPP
enters the pathway after completion of the ring structure.

4. CTP is formed by the amination of UTP.

5. Deoxyribonucleotides are synthesized by the reduction of
ribonucleotides at C-2' in a reaction catalyzed by ribonucleotide
reductase.

6. Thymidylate (dTMP) is formed from deoxyuridylate (dUMP) by a
methylation reaction in which 5,10-methylenetetrahydrofolate

donates both a one-carbon group and a hydride ion. 7,8-Dihydrofo-
late, the other product of this methylation, is recycled by NADPH-
dependent reduction to the active coenzyme tetrahydrofolate.

7. PRPP reacts with pyrimidines and purines in salvage reactions to
yield nucleoside monophosphates. Nucleotides and their con-
stituents are interconverted by a variety of enzymes.

8. Nitrogen from amino acids and purine nucleotides is excreted as
uric acid in birds and some reptiles. Primates degrade purines to
uric acid (urate). Most other organisms further catabolize urate
to allantoin, allantoate, urea, or ammonia.

9. Pyrimidines are catabolized to ammonia, bicarbonate, and either
acetyl CoA (from cytosine or uracil) or succinyl CoA (from
thymine) .

Problems

1. Indicate where the label appears in the product for each of the fol-
lowing precursor-product pairs:

(a) 15 N-aspartate — » AMP

(b) 2- [ 14 C] -glycine — » AMP

(d) 2- [ 14 C] -aspartate — > UMP

(e) H 14 CO 3 0 UMP

2. How many ATP equivalents are needed to synthesize one mole-
cule of IMP, starting from ribose 5 -phosphate? Assume that all
necessary precursors in the pathway are present.

3. The incorporation of one-carbon units in the de novo pathways of
purines and pyrimidines requires tetrahydrofolate (THF) deriva-
tives as donors. List the reactions requiring THF derivatives, indi-
cate the THF donor, and indicate which carbon of the purine or
pyrimidine is derived from THF.

4. The glutamine analog acivicin, a potential
anticancer agent, slows the rapid growth of
cells by inhibiting nucleotide biosynthesis.

(a) Show how acivicin structurally resem-
bles glutamine.

(b) What intermediate accumulates in the
purine biosynthetic pathway when
acivicin is present?

5. A hypothetical bacterium synthesizes UMP by a pathway analo-
gous to the pathway in E. coli , except that /3- alanine is used instead
of aspartate.

/3-Alanine

(a) Why would this pathway be shorter than the pathway in
E. coli ?

(b) When /3-alanine uniformly labeled with 14 C is used, where
would the label appear in UMP?

6. (a) The enzyme dCMP deaminase can provide a major route

from cytidine to uridine nucleotides. What is the product of
the action of dCMP deaminase on dCMP?

(b) This allosteric enzyme is subject to inhibition by dTTP and
activation by dCTP. Explain why this is reasonable in terms of
the overall cellular needs of nucleoside triphosphates.

7. In eukaryotes, how many ATP equivalents are needed to synthe-
size one molecule of UMP from HC0 3 ®, aspartate, glutamine,
and ribose 5-phosphate? (Ignore any ATP that might be produced
by oxidizing the QH 2 generated in the pathway.)

8. Severe combined immunodeficiency syndrome (SCIDS) is char-
acterized by the lack of an immune response to infectious dis-
eases. One form of SCIDS is caused by a deficiency of adenosine
deaminase (ADA), an enzyme that catalyzes the deamination of
adenosine and deoxyadenosine to produce inosine and deoxyino-
sine, respectively. The enzyme deficiency increases dATP levels
but decreases the levels of other deoxynucleotides, thereby inhibit-
ing DNA replication and cell division in certain rapidly dividing
cells. Explain how an adenosine deaminase deficiency affects the
levels of deoxynucleotides. (The first effective gene therapy in hu-
mans was carried out by transforming a patient’s T-cells with a
normal ADA gene.)

9. One cause of gout is a deficiency in hypoxanthine-guanine phos-
phoribosyltransferase activity (Box 18.4). Another cause is due to
an increase in PRPP synthetase activity. If PRPP is a positive effec-
tor of glutamine-PRPP amidotransferase in humans, how does
this affect purine synthesis?

10. Identify the nucleotides involved in the following pathways:

(a) the nucleoside triphosphate required as a substrate in the
synthesis of NAD

(b) the nucleoside triphosphate required in the synthesis of
FMN

(d) the substrate for G proteins

(e) the nucleotide used in the synthesis of glycogen from glucose
6-phosphate

(f) the cofactor required in the reaction catalyzed by mam-
malian succinyl- CoA synthetase

(g) the cosubstrate required for the synthesis of phosphatidylser-
ine from phosphatidate

coa

H 3 N-

-C — H

cA x ch 2

\ /

N=C

Acivicin

572 CHAPTER 18 Nucleotide Metabolism

(h) the nucleotide required for activation of galactose in cerebro-
side biosynthesis

(i) the nucleotide substrate used in histidine biosynthesis

(j) the common precursor of AMP and GMP

(k) the precursor of hypoxanthine

11. The catabolism of fats and carbohydrates provides considerable
metabolic energy in the form of ATP. Does the degradation of
purines and pyrimidines provide a significant source of energy in
eukaryotic cells?

12. PPRP synthetase uses ct-D-ribose 5-phosphate as a substrate. How
is the a isomer formed inside the cell?

13. The systematic names of the common bases are given in Sections
18.1 and 18.2. What are the systematic names of xanthine, hypox-
anthine, and orotate?

14. The sequential action of adenylosuccinate synthetase and adeny-
losuccinate lyase results in the transfer of an amino group from
aspartate and the release of fumarate. Identify two other pairs of
enzymes that accomplish the same goal.

Selected Readings

Purine Metabolism

Honzatko, R. B., Stayton, M. M., and Fromm, H. J.
(1999). Adenylosuccinate synthetase: recent devel-
opments. Adv. Enzymol. Relat. Areas Mol. Biol
73:57-102.

Cendron, L., Berni, R., Folli, C., Ramazzina, I.,
Percudani, R., and Zanotti, G. (2007). The structure
of 2 - oxo - 4 -hydroxy-4 - carb oxy- 5 - ureidoimidazo -
line decarboxylase provides insights into the
mechanism of uric acid degradation. / Biol. Chem.
282:18182-18189.

Kresge, N., Simoni, R. D., and Hill, R. L. (2006).
Biosynthesis of purines: the work of John M.
Buchanan./. Biol. Chem. 281:e35-e36.

Ramazzina, I., Folli, C., Secchi, A., Berni, R., and
Percudani, R. (2006). Completing the uric acid
degradation pathway through phylogenetic com-
parison of whole genomes. Nat Chem Biol.
2:144-148.

Tipton, P. A. (2006). Urate to allantoin, specifically
(S)-allantoin. Nat. Chem. Biol. 2:124-125.

Tsai. M., Koo, J., Yip, P., Colman, R. F., Segall,

M. L., Howell, P. L. (2007). Substrate and product
complexes of Escherichia coli adenylosuccinate
lyase provide new insights into the enzymatic
mechanism./. Mol. Biol. 370:541-554.

Zhang, R.-G., Evans, G., Rotella, F. J., Westbrook,

E. M., Beno, D., Huberman, E., Joachimiak, A., and

Collart, F. R. (1999). Characteristics and crystal
structure of bacterial inosine-5' -monophosphate
dehydrogenase. Biochem. 38:4691-4700.

Pyrimidine Metabolism

Blakley, R. L. (1995). Eukaryotic dihydrofolate
reductase. Adv. Enzymol. Relat. Areas Mol. Biol.
70:23-102.

Carreras, C. W., and Santi, D. V. (1995). The cat-
alytic mechanism and structure of thymidylate
synthase. Ann u. Rev. Biochem. 64:721-762.

Chan, R. S., Sakash, J. B., Macol, C. P., West, J. M.,
Tsuruta, H., and Kantrowitz, E. R. (2002). The role
of intersubunit interactions for the stabilization of
the T state of Escherichia coli aspartate transcar-
bamoylase./. Biol. Chem. 277:49755-49760.

Lipscomb, W. N. (1994). Aspartate transcarbamoy-
lase from Escherichia coli: activity and regulation.
Adv. Enzymol. Relat. Areas Mol. Biol. 68:67-151.

Raushel, F. M., Thoden, J. B., and Holden, H. M.
(1999). The amidotransferase family of enzymes:
molecular machines for the production and deliv-
ery of ammonia. Biochem. 38:7891-7899.

Stroud, R. M. (1994). An electrostatic highway.
Struct. Biol. 1:131-134.

Ribonucleotide Reduction

Eriksson, M., Uhlin, U., Ramaswamy, S., Ekberg,
M., Regnstrom, K., Sjoberg, B. M., and Eklund, H.

(1997). Binding of allosteric effectors to ribonu-
cleotide reductase protein Rl: reduction of active-
site cysteines promotes substrate binding. Structure
5:1077-1092.

Gorlatov, S. N., and Stadtman, T. C. (1998).
Human thioredoxin reductase from HeLa cells:
selective alkylation of selenocysteine in the protein
inhibits enzyme activity and reduction with
NADPH influences affinity to heparin. Proc. Natl.
Acad. Sci. USA. 95:8520-8525.

Jordan, A., and Reichard, P. (1998). Ribonu-
cleotide reductases. Annu. Rev. Biochem. 67:71-98.

Kresge, N., Simoni, R. D., and Hill, R. L. (2006).
Peter Reichard and the reduction of ribonucleo-
sides. /. Biol. Chem. 281:el3-el5.

Nordland, P. and Reichard, P. (2006). Ribonu-
cleotide reductases. Annu. Rev. Biochem.
75:681-706.

Sjoberg, B.M. (2010). A never-ending story.

Science 329:1475-1476.

Stubbe, J. (1998). Ribonucleotide reductases in the
twenty- first century. Proc. Natl. Acad. Sci. USA.
95:2723-2724.

Uppsten, M., Farnegardh, M., Domkin, V., and
Uhlin, U. (2006). The first holocomplex structure
of ribonucleotide reductase gives new insight into
its mechanism of action. /. Mol. Biol. 359:365-377.

o c

_ o

° o o o

° o

o o

° c

o o

Nucleic Acids

T he discovery of the substance that proved to be deoxyribonucleic acid (DNA)
was made in 1869 by Friedrich Miescher, a young Swiss physician working in
the laboratory of the German physiological chemist Felix Hoppe-Seyler. Miescher
treated white blood cells (which came from the pus on discarded surgical bandages)
with hydrochloric acid to obtain nuclei for study. When the nuclei were subsequently
treated with acid, a precipitate formed that contained carbon, hydrogen, oxygen, nitro-
gen, and a high percentage of phosphorus. Miescher called the precipitate “nuclein” be-
cause it came from nuclei. Later, when it was found to be strongly acidic, its name was
changed to nucleic acid. Although he did not know it, Miescher had discovered DNA.
Soon afterward, Hoppe-Seyler isolated a similar substance from yeast cells — this sub-
stance is now known to be ribonucleic acid (RNA). Both DNA and RNA are polymers
of nucleotides, or polynucleotides.

In 1944 Oswald Avery, Colin MacLeod, and Maclyn McCarty demonstrated that
DNA is the molecule that carries genetic information. At the time, very little was known
about the three-dimensional structure of this important molecule. Over the next few
years, the structures of nucleotides were determined and in 1953 James D. Watson and
Francis H. C. Crick proposed their model for the structure of double- stranded DNA.

The study of nucleic acid biochemistry has advanced considerably in the past few
decades. Today it is possible not only to determine the sequence of your genome but
also to synthesize large chromosomes in the laboratory. It has become routine to clone
and manipulate DNA molecules. This has led to spectacular advances in our under-
standing of molecular biology and the ways information contained in DNA is expressed
in living cells.

We now know that a living organism contains a set of instructions for every step re-
quired to construct a replica of itself. This information resides in the genetic material,
or genome, of the organism. The genomes of all cells are composed of DNA but some
viral genomes are composed of RNA. A genome may consist of a single molecule of
DNA, as in many species of bacteria. The genome of eukaryotes is one complete set
of DNA molecules found in the nucleus (i.e., the haploid set of chromosomes in diploid
organisms). By convention, the genome of a species does not include mitochondrial
and chloroplast DNA. With rare exception, no two individuals in a species have exactly

We wish to suggest a structure for
the salt of deoxyribose nucleic acid
(D.N.A.). This structure has novel
features which are of considerable
biological interest.

— J.D. Watson and F.H.C. Crick (1953)

Top: Space-filling model of DNA, viewed along the helix axis.

573

574 CHAPTER 19 Nucleic Acids

▲ James D. Watson (1928-) (left) and Francis
H. C. Crick (1916-2004) (right) describing
the structure of DNA in 1953.

The distinction between the normal
flow of information and the Central
Dogma of Molecular Biology is
explained in Section 1.1 and the intro-
duction to Chapter 21.

▲ Figure 19.1

Chemical structure of a nucleotide. Nucleotides
contain a five-carbon sugar, a nitrogenous
base, and at least one phosphate group. The
sugar can be either deoxyribose (shown here)
or ribose.

HOCH 2 x 0n OH

Ribose

(/3-D-Ribofuranose)

Deoxyribose

(2-Deoxy-/3-D-ribofuranose)

▲ Figure 19.2

Chemical structures of the two sugars found in
nucleotides, (a) Ribose (/kD-ribofuranose).
(b) Deoxyribose (2-deoxy-/3-D-ribofuranose).

the same genome sequence. If they were alive today, Miescher and Hoppe-Seyler would
be astonished to learn that criminals could be convicted by DNA fingerprinting and
that we have sequenced the complete genomes of thousands of species, including
humans.

In general, the information that specifies the primary structure of a protein is en-
coded in the sequence of nucleotides in DNA. This information is enzymatically copied
during the synthesis of RNA, a process known as transcription. Some of the informa-
tion contained in the transcribed RNA molecules is translated during the synthesis of
polypeptide chains that are then folded and assembled to form protein molecules. Thus,
we can generalize that the biological information stored in a cell’s DNA flows from
DNA to RNA to protein.

Nucleic acids are the fourth major class of macromolecules that we study in this
book. Like proteins and polysaccharides, they contain multiple similar monomeric
units that are covalently joined to produce large polymers. In this chapter we describe
the structure of nucleic acids and how they are packaged in cells. We also describe some
of the enzymes that use DNA and RNA as substrates. Many other proteins and enzymes
interact with DNA and RNA in order to ensure that genetic information is correctly in-
terpreted. We will consider the biochemistry and the regulation of this flow of informa-
tion in Chapters 20 to 22.

19.1 Nucleotides Are the Building
Blocks of Nucleic Acids

Nucleic acids are polynucleotides, or polymers of nucleotides. As we saw in the previous
chapter, nucleotides have three components: a five-carbon sugar, one or more phos-
phate groups, and a weakly basic nitrogenous compound called a base (Figure 19.1).
The bases found in nucleotides are substituted pyrimidines and purines. The pentose is
either ribose (D-ribofuranose) or 2-deoxyribose (2-deoxy-D-ribofuranose). The pyrim-
idine or purine N-glycosides of these sugars are called nucleosides. Nucleotides are the
phosphate esters of nucleosides — the common nucleotides contain from one to three
phosphoryl groups. Nucleotides containing ribose are called ribonucleotides and nu-
cleotides containing deoxyribose are called deoxyribonucleo tides (Section 18.5).

A. Ribose and Deoxyribose

The sugar components of the nucleotides found in nucleic acids are shown in Figure 19.2.
Both sugars are shown as Haworth projections of the /3- conformation of the furanose
ring forms (Section 8.2). This is the stable conformation found in nucleotides and
polynucleotides. Each of these furanose rings can adopt different conformations such as
the envelope forms discussed in Chapter 8. The 2'-endo conformation of deoxyribose
predominates in double-stranded DNA (Figure 8.11).

B. Purines and Pyrimidines

The bases found in nucleotides are derivatives of either pyrimidine or purine (Chapter 18).
The structures of these heterocyclic compounds and the numbering systems for the car-
bon and nitrogen atoms of each base are shown in Figure 19.3. Pyrimidine has a single
ring containing four carbon and two nitrogen atoms. Purine has a fused pyrimidine-
imidazole ring system. Both types of bases are unsaturated, with conjugated double
bonds. This feature makes the rings planar and also accounts for their ability to absorb
ultraviolet light.

Substituted purines and pyrimidines are ubiquitous in living cells but the unsubsti-
tuted bases are seldom encountered in biological systems. The major pyrimidines that
occur in nucleotides are uracil (2,4-dioxopyrimidine, U), thymine (2,4-dioxo-5-
methylpyrimidine, T), and cytosine (2-oxo-4-aminopyrimidine, C). The major purines
are adenine (6-aminopurine, A) and guanine (2-amino-6-oxopurine, G). The chemical
structures of these five major bases are shown in Figure 19.4. Note that thymine can
also be called 5-methyluracil because it is a substituted form of uracil (Section 18.6).

19.1 Nucleotides Are the Building Blocks of Nucleic Acids 575

Adenine, guanine, and cytosine are found in both ribonucleotides and deoxyribonu-
cleotides. Uracil is found mainly in ribonucleotides and thymine is found mainly in de-
oxyribonucleotides.

Purines and pyrimidines are weak bases and are relatively insoluble in water at
physiological pH. Within cells, however, most pyrimidine and purine bases occur as con-
stituents of nucleotides and polynucleotides and these compounds are highly soluble.

Each heterocyclic base can exist in at least two tautomeric forms. Adenine and cyto-
sine can exist in either amino or imino forms. Guanine, thymine, and uracil can exist in
either lactam (keto) or lactim (enol) forms (Figure 19.5). The tautomeric forms of each
base exist in equilibrium but the amino and lactam tautomers are more stable and
therefore predominate under the conditions found inside most cells. Note that the rings
remain unsaturated and planar in each tautomer.

All of the bases in the common nucleotides can participate in hydrogen bonding.
The amino groups of adenine and cytosine are hydrogen donors and the ring nitrogen
atoms (N-l in adenine and N-3 in cytosine) are hydrogen acceptors (Figure 19.6). Cyto-
sine also has a hydrogen acceptor group at C-2. Guanine, cytosine, and thymine can form
three hydrogen bonds. In guanine, the group at C-6 is a hydrogen acceptor while N-l and
the amino group at C-2 are hydrogen donors. In thymine, the groups at C-4 and C-2 are
hydrogen acceptors and N-3 is a hydrogen donor. (Only two of these sites, C-4 and N-3,
are used to form base pairs in DNA.) The hydrogen-bonding ability of uracil, a base
found in RNA, is similar to that of thymine. The hydrogen-bonding patterns of bases
have important consequences for the three-dimensional structure of nucleic acids.

Biochemistry textbooks in the 1940s usually depicted the bases in their imino and
lactim forms. These were the structures that Jim Watson was using in 1953 to build a
model of DNA. Shortly after being told by Jerry Donohue that the textbooks were
wrong, Watson discovered the now- famous A/T and G/C base pairs.

Additional hydrogen bonding occurs in some nucleic acids and in nucleic
acid-protein interactions. For example, N-7 of adenine and guanine can be a hydrogen
acceptor and both amino hydrogen atoms of adenine, guanine, and cytosine can be do-
nated to form hydrogen bonds.

C. Nucleosides

Nucleosides are composed of ribose or deoxyribose and a heterocyclic base. In each nu-
cleoside, a /3-N-glycosidic bond connects C-l of the sugar to N-l of the pyrimidine or
N-9 of the purine. Nucleosides are therefore N-ribosyl or N-deoxyribosyl derivatives of
pyrimidines or purines. The numbering convention for carbon and nitrogen atoms in

PYRIMIDINES

H H H

Uracil Thymine Cytosine

(2,4-Dioxopyrimidine) (2,4-Dioxo-5-methylpyrimidine) (2-Oxo-4-aminopyrimidine)

PURINES

NH 2 O

Adenine Guanine

(6-Aminopurine) (2-Amino-6-oxopurine)

Purine

▲ Figure 19.3

Chemical structures of pyrimidine and purine.

◄ Figure 19.4

Chemical structures of the major pyrimidines
and purines.

576 CHAPTER 19 Nucleic Acids

Figure 19.5 ►

Tautomers of adenine, cytosine, guanine,
thymine, and uracil. At physiological pH, the
equilibria of these tautomerization reactions
lie far in the direction of the amino and lac-
tam forms.

Adenine

Cytosine

Guanine

Thymine

Uracil

Predominant forms

Lactam Lactim

(Deoxy)Adenosine

(Deoxy)Cytidine

(Deoxy)Thymidine

nucleosides reflects the fact that they are composed of a base and a
five-carbon sugar, each of which has its own numbering scheme.
The designation of atoms in the purine or pyrimidine moieties takes
precedence. Hence the atoms in the bases are numbered 1, 2, 3, and so
on, while those in the furanose ring are distinguished by adding primes
('). Thus, the /3-N-glycosidic bond connects the C-l', or 1', atom of
the sugar moiety to the base. Ribose and deoxyribose differ at the
C-2', or 2', position. The chemical structures of the major ribonucle-
osides and deoxyribonucleosides are shown in Figure 19.7.

The names of nucleosides are derived from the names of their
bases. The ribonucleoside containing adenine is called adenosine
(the systematic name, 9-/3-D-ribofuranosyladenine, is seldom used)

◄ Figure 19.6

Hydrogen bond sites of bases in nucleic acids. Each base contains atoms and
functional groups that can serve as hydrogen donors or acceptors. The com-
mon tautomeric forms of the bases are shown. Hydrogen donor and acceptor
groups differ in the other tautomers. R represents the sugar moiety.

19.1 Nucleotides Are the Building Blocks of Nucleic Acids 577

Adenosine

Guanosine

Cytidine

Uridine

Deoxyadenosine Deoxyguanosine Deoxycytidine Deoxythymidine

(Thymidine)

and its deoxy counterpart is called deoxyadenosine. Similarly, the ribonucleosides of
guanine, cytosine, and uracil are guanosine, cytidine, and uridine, respectively. The deoxyri-
bonucleosides of guanine, cytosine, and thymine are deoxyguanosine, deoxycytidine,
and deoxythymidine, respectively. Deoxythymidine is often simply called thymidine be-
cause thymine rarely occurs in ribonucleosides. The single-letter abbreviations for
pyrimidine and purine bases are also commonly used to designate ribonucleosides:
A, G, C, and U (for adenosine, guanosine, cytidine, and uridine, respectively). The de-
oxyribonucleosides are abbreviated dA, dG, dC, and dT when it is necessary to distin-
guish them from ribonucleosides.

Rotation around the glycosidic bonds of nucleosides and nucleotides is sometimes
hindered. There are two relatively stable conformations, syn and anti , that are in rapid
equilibrium (Figure 19.8). In the common pyrimidine nucleosides, the anti conforma-
tion predominates. The anti conformations of all nucleotides predominate in nucleic
acids, the polymers of nucleotides.

▲ Figure 19.7

Chemical structures of nucleosides.

Note that the carbon atoms of the sugars are
numbered with primes to distinguish them
from the atoms of the bases, (a) Ribonucleo-
sides. The sugar in ribonucleosides is
ribose, which contains a hydroxyl group at
C-2\ as shown here. The /TA/-glycosidic
bond of adenosine is shown in red.

(b) Deoxyribonucleosides. In deoxyribonucle-
osides, there is an additional hydrogen atom
at C-2' instead of a hydroxyl group.

D. Nucleotides

Nucleotides are phosphorylated derivatives of nucleosides. Ribonucleosides contain
three hydroxyl groups that can be phosphorylated (2', 3', and 5'), and deoxyribonucle-
osides contain two such hydroxyl groups (3' and 5'). The phosphoryl groups in natu-
rally occurring nucleotides are usually attached to the oxygen atom of the 5 '-hydroxyl
group. By convention, a nucleotide is always assumed to be a 5 '-phosphate ester unless
otherwise designated.

The systematic names for nucleotides indicate the number of phosphate groups
present. For example, the 5 '-monophosphate ester of adenosine is known as adenosine
monophosphate (AMP). It is also simply called adenylate. Similarly, the 5 '-monophosphate
ester of deoxycytidine can be referred to as deoxycytidine monophosphate (dCMP) or
deoxycytidylate. The 5 ' -monophosphate ester of the deoxyribonucleoside of thymine is
usually known as thymidylate but is sometimes called deoxythymidylate to avoid ambi-
guity. Table 19.1 presents an overview of the nomenclature of bases, nucleosides, and
5 '-nucleotides. Nucleotides with the phosphate esterified at the 5' position are abbrevi-
ated AMP, dCMP, and so on. Nucleotides with the phosphate esterified at a position
other than 5' are given similar abbreviations but with position numbers designated
(e.g., 3 '-AMP).

KEY CONCEPT

By convention the numbering of the atoms
in the base takes precedence so the
carbon atoms in the sugar are numbered
1 ' (“one prime”), 2' (“two prime”), etc.

578 CHAPTER 19 Nucleic Acids

OH OH

syn Adenosine

NH 2

OH OH

anti Adenosine
▲ Figure 19.8

Syn and anti conformations of adenosine.

Some nucleosides assume either the syn or
anti conformation. The anti form is usually
more stable in pyrimidine nucleosides.

Figure 19.9 ►

Chemical structures of the deoxyribonucleo-
side-5 '-monophosphates.

Table 19.1 Nomenclature of bases, nucleosides, and nucleotides

Base

Ribonucleoside

Ribonucleotide
(5 '-monophosphate)

Adenine (A)

Adenosine

Adenosine 5'-monophoshate (AMP); adenylate 9

Guanine (G)

Guanosine

Guanosine 5'-monophosphate (GMP); guanylate 9

Cytosine (C)

Cytidine

Cytidine 5'-monophosphate (CMP); cytidylate 9

Uracil (U)

Uridine

Uridine 5 '-monophosphate (UMP); uridylate 9

Base

Deoxyribonudeoside

Deoxyribonudeotide
(5 '-monophosphate)

Adenine (A)

Deoxyadenosine

Deoxyadenosine 5 '-monophosphate (dAMP);
deoxyadenylate 9

Guanine (G)

Deoxyguanosine

Deoxyguanosine 5 '-monophosphate (dGMP);
deoxyguanylate 9

Cytosine (C)

Deoxycytidine

Deoxycytidine 5'-monophosphate (dCMP);
deoxycytidylate 9

Thymine (T)

Deoxythymidine
or thymidine

Deoxythymidine 5 '-monophosphate (dTMP);
deoxythymidylate 9 or thymidylate 9

a Anionic forms of phosphate esters predominant at pH 7.4.

Nucleoside monophosphates, which are derivatives of phosphoric acid, are anionic at
physiological pH. They are dibasic acids under physiological conditions since the piC a values
are approximately 1 and 6. The nitrogen atoms of the heterocyclic rings can also ionize.

Nucleoside monophosphates can be further phosphorylated to form nucleoside
diphosphates and nucleoside triphosphates. These additional phosphoryl groups are
present as phosphoanhydrides. The chemical structures of the deoxyribonucleoside-5'-
monophosphates are shown in Figure 19.9. A three-dimensional view of the structure

0 °

2'-Deoxyadenosine 5'-monophosphate
(Deoxyadenylate f dAMP)

0 °

2'-Deoxyguanosine 5'-monophosphate
(Deoxyguanylate, dGMP)

O 0

O 1

°0— P = 0 o

OH H

2'-Deoxycytidine 5'-monophosphate
(Deoxycytidylate, dCMP)

2'-Deoxythymidine 5'-monophosphate
(Thymidylate, dTMP)

19.2 DNA Is Double-Stranded 579

of dGMP is shown in Figure 19.10. The base in dGMP is in the anti conformation and
the sugar ring is puckered. The plane of the purine ring is almost perpendicular to that
of the furanose ring. The phosphoryl group attached to the 5 '-carbon atom is posi-
tioned well above the sugar and far away from the base.

Nucleoside polyphosphates and polymers of nucleotides can also be abbreviated
using a scheme in which the phosphate groups are represented by a p” and the nucleo-
sides are represented by their one-letter abbreviations. The position of the a p” relative to
the nucleoside abbreviation indicates the position of the phosphate — for a 5' phos-
phate, the p precedes the nucleoside abbreviation and for a 3 ' phosphate, the a p” follows
the nucleoside abbreviation. Thus, 5 '-adenylate (AMP) can be abbreviated as pA, 3'-
deoxyadenylate as dAp, and ATP as pppA.

▲ Figure 19.10

Deoxyguanosine-5'-monophosphate (dGMP).

Hydrogen atoms have been omitted for clarity.
Color key: carbon, black; nitrogen, blue; oxygen,
red; phosphorus, purple.

19.2 DNA Is Double-Stranded

By 1950 it was clear that DNA is a linear polymer of 2'-deoxyribonucleotide residues
linked by 3 '-5' phosphodiesters. Moreover, Erwin Chargaff had deduced certain regu-
larities in the nucleotide compositions of DNA samples obtained from a wide variety of
prokaryotes and eukaryotes. Among other things, Chargaff observed that in the DNA
of a given cell, A and T are present in equimolar amounts, as are G and C. An example
of modern DNA composition data showing these ratios is presented in Table 19.2.
Although A = T and G = C for each species, the total mole percent of (G + C) may dif-
fer considerably from that of (A + T). The DNA of some organisms, such as the yeast
Saccharomyces cerevisiae , is relatively deficient in (G + C) whereas the DNA of other or-
ganisms, such as the bacterium Mycobacterium tuberculosis , is rich in (G + C). In gen-
eral, the DNAs of closely related species, such as cows, pigs, and humans, have similar
base compositions. The data also shows that the ratio of purines to pyrimidines is al-
ways 1:1 in the DNA of all species.

The model of DNA proposed by Watson and Crick in 1953 was based on the
known structures of the nucleosides and on X-ray diffraction patterns that Rosalind
Franklin and Maurice Wilkins obtained from DNA fibers. The Watson-Crick model
accounted for the equal amounts of purines and pyrimidines by suggesting that DNA
was double- stranded and that bases on one strand paired specifically with bases on the
other strand: A with T and G with C. Watson and Crick’s proposed structure is now re-
ferred to as the B conformation of DNA, or simply B-DNA.

An appreciation of DNA structure is important for understanding the processes of
DNA replication (Chapter 20) and transcription (Chapter 21). DNA is the storehouse
of biological information. Every cell contains dozens of enzymes and proteins that bind
to DNA recognizing certain structural features, such as the sequence of nucleotides. In
the following sections we will see how the structure of DNA allows these proteins to
gain access to the stored information.

Table 19.2 Base composition of DNA (mole %) and ratios of bases

Source

A/T"

c/c°

(C + C)

Purine/

pyrimidine"

Escherichia coli

26.0

24.9

25.2

23.9

1.09

0.99

50.1

1.04

Mycobacterium tuberculosis

15.1

34.9

35.4

14.6

1.03

0.99

70.3

1.00

Yeast

31.7

18.3

17.4

32.6

0.97

1.05

35.7

1.00

Cow

29.0

21.2

28.7

1.01

1.00

42.4

1.01

Pig

29.8

20.7

29.1

1.02

1.00

41.4

1.01

Human

30.4

19.9

30.1

1.01

1.00

39.8

1.01

deviations from a 1 :1 ratio are due to experimental variations.

580 CHAPTER 19 Nucleic Acids

A linkage group consists of several
different covalent bonds.

Figure 19.11 ►

Chemical structure of the tetranucleotide
pdApdGpdTpdC. The nucleotide residues are
joined by 3'-5' phosphodiester linkages.
The nucleotide with a free 5'-phosphoryl
group is called the 5' end, and the
nucleotide with a free 3'-hydroxyl group
is called the 3' end.

A. Nucleotides Are Joined by 3'-5' Phosphodiester Linkages

We have seen that the primary structure of a protein refers to the sequence of its amino acid
residues linked by peptide bonds. Similarly, the primary structure of a nucleic acid is the
sequence of its nucleotide residues connected by 3 '-5' phosphodiester linkages. A tetranu-
cleotide representing a segment of a DNA chain illustrates such linkages (Figure 19.11).
The backbone of the polynucleotide chain consists of the phosphoryl groups, the 5', 4',
and 3' carbon atoms, and the 3' oxygen atom of each deoxyribose. These backbone
atoms are arranged in an extended conformation. This makes double-stranded DNA a
long, thin molecule, unlike polypeptide chains that can easily fold back on themselves.

All the nucleotide residues within a polynucleotide chain have the same orienta-
tion. Thus, polynucleotide chains have directionality, like polypeptide chains. One end
of a linear polynucleotide chain is said to be 5' (because no residue is attached to its 5'-
carbon) and the other is said to be 3' (because no residue is attached to its 3 '-carbon).
By convention, the direction of a DNA strand is defined by reading across the atoms
that make up the sugar residue. Thus, going from the top to the bottom of the strand in

5' end

u o— p=o

5 ' CH ^ o. N— n ;

4 '

nh 2

< 7 J J

Adenine (A)

3'-5'
phosphodiester -<
linkage

H H

O H

o — P = o

5'CH 2

4 '

Guanine (G)

o^n^n^nh.

3'-5'
phosphodiester «
linkage

H H

O H

u o— p = o

5'CH 2

6 h bC.

Ov N

Thymine (T)

3'-5'
phosphodiester -<
linkage

O H

°o — P = o

NH,

,(Y

Cytosine (C)

OH H

3' end

2 '

19.2 DNA Is Double-Stranded 581

Figure 19.1 1 is defined as 5' —> 3' (“five prime to three prime”) because one crosses the
sugar residue encountering the 5', 4', and 3' carbon atoms, in that order. Similarly,
going from the bottom to the top of the strand is moving in the 3' — > 5' direction.

Structural abbreviations are assumed to read in the 5' —> 3' direction unless other-
wise specified. Because phosphates can be abbreviated as “p,” the tetranucleotide in
Figure 19.11 can be referred to as pdApdGpdTpdC, or shortened to AGTC when it is
clear that the reference is to DNA.

Each phosphate group that participates in a phosphodiester linkage has a pK a of
about 2 and bears a negative charge at neutral pH. Consequently, nucleic acids are
polyanions under physiological conditions. Negatively charged phosphate groups are
neutralized by small cations and positively charged proteins.

B. Two Antiparallel Strands Form a Double Helix

Most DNA molecules consist of two strands of polynucleotide. Each of the bases on one
strand forms hydrogen bonds with a base of the opposite strand (Figure 19.12). The

KEY CONCEPT

The direction of moving along a DNA
or RNA strand can be either 5' —> 3' or
3' —> 5'. It is defined by the direction
of reading across the atoms that make up
the sugar residue.

▲ Watson and Crick’s original DNA model.

◄ Figure 19.12

Chemical structure of double-stranded DNA.

The two strands run in opposite directions.
Adenine in one strand pairs with thymine in
the opposite strand, and guanine pairs with
cytosine.

5 '

582

CHAPTER 19 Nucleic Acids

KEY CONCEPT

The two strands of DNA are anti-parallel.

most common base pairs occur between the lactam and amino tautomers of the bases.
Guanine pairs with cytosine and adenine with thymine in a manner that maximizes
hydrogen bonding between potential sites. G/C base pairs have three hydrogen bonds
and A/T base pairs have two. This feature of double-stranded DNA accounts for Char-
gaff’s discovery that the ratio of A to T and of G to C is 1:1 for a wide variety of DNA
molecules. Because A in one strand pairs with T in the other strand and G pairs with C,
the strands are complementary and each one can serve as a template for the other.

The sugar-phosphate backbones of the complementary strands of double-
stranded DNA have opposite orientations. In other words, they are antiparallel. This
was one of the important new insights contributed by Watson and Crick when they
built their model of DNA in 1953.

Each end of double-stranded DNA is made up of the 5' end of one strand and the
3' end of another. The distance between the two sugar-phosphate backbones is the
same for each base pair. Consequently, all DNA molecules have the same regular struc-
ture in spite of the fact that their nucleotide sequences may be quite different.

The actual structure of DNA differs in two important aspects from that shown in
Figure 19.12. In a true three-dimensional representation, the two strands wrap around
each other to form a two-stranded helical structure, or double helix. Also, the bases are
rotated so that the plane of the base pairs is nearly perpendicular to the page. (Recall
that the plane of the base in dGMP is nearly perpendicular to that of the sugar, as
shown in Figure 19.10.)

The DNA molecule can be visualized as a “ladder” that has been twisted into a
helix. The paired bases represent the rungs of the ladder and the sugar-phosphate back-
bones represent the supports. Each complementary strand serves as a perfect template
for the other. This complementarity is responsible for the overall regularity of the struc-
ture of double-stranded DNA. However, complementary base pairing alone does not
produce a helix. In B-DNA, the base pairs are stacked one above the other and are nearly
perpendicular to the long axis of the molecule. The cooperative, noncovalent interactions
between the upper and lower surfaces of each base pair bring the pairs closer together
and create a hydrophobic interior that causes the sugar-phosphate backbone to twist. It
is these stacking interactions that create the familiar helix (Figure 19.13). Much of the
stability of double-stranded DNA is due to the stacking interactions between base pairs.

The two hydrophilic sugar-phosphate backbones wind around the outside of the
helix where they are exposed to the aqueous environment. In contrast, the stacked, rela-
tively hydrophobic bases are located in the interior of the helix where they are largely
inaccessible to water. This hydrophobic environment makes the hydrogen bonds be-
tween bases more stable since they are shielded from competition with water molecules.

The double helix has two grooves of unequal width because of the way the base
pairs stack and the sugar-phosphate backbones twist. These grooves are called the major
groove and the minor groove (Figure 19.14). Within each groove, functional groups on
the edges of the base pairs are exposed to water. Each base pair has a distinctive pattern
of chemical groups projecting into the grooves. Molecules that interact with particular
base pairs can identify them by binding in the grooves without disrupting the helix.
This is particularly important for proteins that must bind to double-stranded DNA and
“read” a specific sequence.

B-DNA is a right-handed helix with a diameter of 2.37 nm. The rise of the helix
(the distance between one base pair and the next along the helical axis) averages 0.33 nm,
and the pitch of the helix (the distance to complete one turn) is about 3.40 nm. These
values vary to some extent depending on the base composition. Because there are about
10.4 base pairs per turn of the helix, the angle of rotation between adjacent nucleotides
within each strand is about 34.6° (360/10.4).

Two views of B-DNA are shown in Figure 19.15. The ball-and-stick model
(Figure 19.15a) shows that the hydrogen bonds between base pairs are buried in the
interior of the molecule where they are protected from competing interactions with water.
The charged phosphate groups (purple and red atoms) are located on the outside surface.
This arrangement is more evident in the space-filling model (Figure 19.15b). The space-
filling model also clearly shows that functional groups of the base pairs are exposed in

19.2 DNA Is Double-Stranded 583

3' 5'

◄ Figure 19.13

Complementary base pairing and stacking
in double-stranded DNA.

2.37 nm

the grooves. These groups can be identified by the presence of blue nitrogen atoms and
red oxygen atoms.

The length of double-stranded DNA molecules is often expressed in terms of base
pairs (bp). For convenience, longer structures are measured in thousands of base pairs,
or kilobase pairs, commonly abbreviated kb. Most bacterial genomes consist of a single
DNA molecule of several thousand kb; for example, the Escherichia coli chromosome is
4600 kb in length. The largest DNA molecules in the chromosomes of mammals and
flowering plants may be several hundred thousand kb long. The human genome con-
tains 3,200,000 kb (3.2 x 10 9 base pairs) of DNA.

Most bacteria have a single chromosome whose ends are joined to create a circular
molecule. DNA in the mitochondria and chloroplasts of eukaryotic cells is also circular.
In contrast, the chromosomes in the nucleus of a eukaryotic cell are linear. (Some bacteria
also have multiple chromosomes and some have linear chromosomes.)

C. Weak Forces Stabilize the Double Helix

The forces that maintain the native conformations of complex cellular structures are
strong enough to maintain the structures but weak enough to allow conformational
flexibility. Covalent bonds between adjacent residues define the primary structures
of proteins and nucleic acids but weak forces determine the three-dimensional shapes

Figure 19.14 ►

Three-dimensional structure of B-DNA. This model shows the orientation of the base pairs and the sugar-
phosphate backbone and the relative sizes of the pyrimidine and purine bases. The sugar-phosphate
backbone winds around the outside of the helix and the bases occupy the interior. Stacking of the
base pairs creates two grooves of unequal width — the major and the minor grooves. The diameter
of the helix is 2.37 nm, and the distance between base pairs is 0.33 nm. The distance to complete
one turn is 3.40 nm. (For clarity, a slight space has been left between the stacked base pairs and
the interactions between complementary bases are shown schematically.)

Minor

groove

Major

groove

Sugar-

phosphate

backbones

Base pair

584 CHAPTER 19 Nucleic Acids

(a)

▲ Figure 19.15

B-DNA. (a) Bal l-and-stick model. The base
pairs are nearly perpendicular to the sugar-
phosphate backbones, (b) Space-filling
model. Color key: carbon, gray; nitrogen,
blue; oxygen, red; phosphorus, purple.
[Nucleic Acids Database BD0001].

of these macromolecules. Four types of interactions affect the conformation of double-
stranded DNA.

1. Stacking interactions. The stacked base pairs form van der Waals contacts.
Although the forces between individual stacked base pairs are weak, they are addi-
tive so in large DNA molecules the van der Waals contacts are an important source
of stability.

2. Hydrogen bonds. Hydrogen bonding between base pairs is a significant stabilizing force.

3. Hydrophobic effects. Burying hydrophobic purine and pyrimidine rings in the inte-
rior of the double helix increases the stability of the helix.

4. Charge-charge interactions. Electrostatic repulsion of the negatively charged phos-
phate groups of the backbone is a potential source of instability in the DNA helix.
However, repulsion is minimized by the presence of cations such as Mg® 1 and
cationic proteins (proteins that contain an abundance of the basic residues arginine
and lysine).

The importance of stacking interactions can be illustrated by examining the vari-
ous stacking energies of the base pairs (Table 19.3). The stacking energy of two base
pairs depends on the nature of the base pair (G/C or A/T) and the orientation of each
base pair. Typical stacking energies are about 35 kj mol" 1 . Within the hydrophobic core
of stacked double-stranded DNA the hydrogen bonds between base pairs have a
strength of about 27 kj mol" 1 each (Section 2.5B). However, if the stacking interac-
tions are weakened, the hydrogen bonds in the base pairs are exposed to competition
from water molecules and the overall contribution to keeping the strands together
diminishes greatly.

Under physiological conditions, double-stranded DNA is thermodynamically
much more stable than the separated strands and that explains why the double-
stranded form predominates in vivo. However, the structure of localized regions of the
double helix can sometimes be disrupted by unwinding. Such disruption occurs during
DNA replication, repair, recombination, and transcription. Complete unwinding and
separation of the complementary single strands is called denaturation. Denaturation
occurs only in vitro.

Double-stranded DNA can be denatured by heat or by a chaotropic agent such
as urea or guanidinium chloride. (Recall from Section 4.10 that proteins can also be
denatured.) In studies of thermal denaturation, the temperature of a solution of DNA is
slowly increased. As the temperature is raised, more and more of the bases become un-
stacked and hydrogen bonds between base pairs are broken. Eventually, the two strands
separate completely. The temperature at which half the DNA has become single-
stranded is known as the melting point (T m ).

Absorption of ultraviolet light can be used to measure the extent of denaturation.
Measurements are made at a wavelength of 260 nm — close to the absorbance maximum
for nucleic acids. Single-stranded DNA absorbs 12% to 40% more light than double-
stranded DNA at 260 nm (Figure 19.16). A plot of the change in absorbance of a DNA
solution versus temperature is called a melting curve (Figure 19.17). The absorbance in-
creases sharply at the melting point and the transition from double-stranded to single-
stranded DNA takes place over a narrow range of temperature.

The sigmoid shape of the melting curve indicates that denaturation is a cooperative
process as we saw in the case of protein denaturation (Section 4.10). In this case, coop-
erativity results from rapid unzippering of the double-stranded molecule as the many
stacking interactions and hydrogen bonds are disrupted. The unzippering begins with
the unwinding of a short internal stretch of DNA, forming a single- stranded “bubble ”
This single-stranded bubble rapidly destabilizes the adjacent stacked base pairs and this
destabilization is propagated in both directions as the bubble expands.

As shown in Figure 19.17, poly (GC) denatures at a much higher temperature than
poly (AT). It is easier to melt A/T-rich DNA than G/C-rich DNA because A/T base pairs
have weaker stacking interactions as shown in Table 19.3. It’s important to note that the
stacking interactions are the first interactions to be disrupted by higher temperature.
Once this process begins the hydrogen bonds — although collectively stronger in stacked

19.2 DNA Is Double-Stranded 585

Figure 19.16 ►

Absorption spectra of double-stranded
and single-stranded DNA. At pH 7.0,
double-stranded DNA has an
absorbance maximum near 260 nm.
Denatured DNA absorbs 12% to
40% more ultraviolet light than
double-stranded DNA.

DNA — become much weaker because they are exposed to water and the DNA is rapidly
destabilized. Naturally occurring DNA is a mixture of regions with varying base com-
positions but A/T-rich regions are more easily unwound than G/C-rich regions.

At temperatures just below the melting point, a typical DNA molecule contains
double-stranded regions that are G/C-rich and local single-stranded regions (“bubbles”)
that are A/T-rich. These in vitro experiments demonstrate an important point — that it
is easier to unwind localized regions whose sequence consists largely of A/T base pairs
rather than G/C base pairs. We will see in Chapter 21 that the initiation sites for tran-
scription are often A/T-rich.

D. Conformations of Double-Stranded DNA

Double-stranded DNA can assume different conformations under different conditions.
X-ray crystallographic studies of various synthetic oligodeoxyribonucleotides of known
sequence indicate that DNA molecules inside the cell do not exist in a “pure” B confor-
mation. Instead, DNA is a dynamic molecule whose exact conformation depends to
some extent on the nucleotide sequence. The local conformation is also affected by
bends in the DNA molecule and whether it is bound to protein. As a result, the number
of base pairs per turn in B-DNA can fluctuate in the range of 10.2-10.6.

There are two other distinctly different DNA conformations in addition to the various
forms of B-DNA. A-DNA forms when DNA is dehydrated and Z-DNA can form when cer-
tain sequences are present (Figure 19.18). (The A- and B-DNA forms were discovered
by Rosalind Franklin in 1952.) A-DNA is more tightly wound than B-DNA and the

Table 19.3 Stacking interactions for the
ten possible combinations in
double-stranded DNA

Stacking

energies

Stacked dimers (kj mol -1 )

'C-G 1 t T-A

A-T I Ig-c

[ - 440

' C-G 1 t A-T 1

T-aI G-cJ

[ - 410

k G-C 1
C-G I

-40.5

^ G-C 1 t C-G 1

G-cl C-gJ

[ ~ 34 - 6

T-A 1
A-T I

-27.5

^ G-C 1 t A-T 1

T-A I 1 C-gJ

[ ~ 27 - 5

v G-C 1 j T-A

t-aI c-gJ

[ ~ 28 - 4

< <
1— 1—

< <

[ - 22 - 5

i si

Arrows designate the direction of the sugar-phos-
phate backbone and point from C-3' of one sugar
unit to C-5' of the next.

[Adapted from Omstein, R. L., Rein, R., Breen, D.
L., and MacElroy, R. D. (1 978). An optimized po-
tential function for the calculation of nucleic acid
interaction energies: I. Base stacking. Biopolymers
17: 2341-2360.]

◄ Figure 19.17 Melting curve for DNA.

In this experiment, the temperature of a DNA so-
lution is increased while the absorbance at 260 nm
is monitored. The melting point (7 m ) corresponds
to the inflection point of the sigmoidal curve
where the increase in absorbance of the sample
is one-half the increase in absorbance of com-
pletely denatured DNA. Poly (AT) melts at a lower
temperature than either naturally occurring DNA
or poly (GC) since more energy is required to
disrupt stacked G/C base pairs.

586 CHAPTER 19 Nucleic Acids

Figure 19.18 ►

A-DNA, B-DNA, and Z-DNA. The A-DNA confor-
mation (left) is favored when DNA is dehy-
drated [NDB AD0001]. B-DNA (center) is
the conformation normally found inside cells
[NDB BDOOOl]. The Z-DNA conformation
(right) is favored in certain G/C-rich
sequences [NDB ZDJ050].

major and minor grooves of A-DNA are similar in width. There are about 1 1 bp per
turn in A-DNA and the base pairs are tilted about 20° relative to the long axis of the
helix. Z-DNA differs even more from B-DNA. There are no grooves in Z-DNA and the
helix is left-handed, not right-handed. The Z-DNA conformation occurs in G/C-rich
regions. Deoxyguanylate residues in Z-DNA have a different sugar conformation
(3'-endo) and the base is in the syn conformation. A-DNA and Z-DNA conformations
exist in vivo but they are confined to short regions of DNA.

v Figure 19.19

Supercoiled DNA. The DNA molecule on the
left is a relaxed closed circle and has the
normal B conformation. Breaking the DNA
helix and unwinding it by two turns before
re-forming the circle produces two super-
coils. The supercoils compensate for the un-
derwinding and restore the normal B confor-
mation. The molecule on the right has a
locally unwound region of DNA. This confor-
mation is topologically equivalent to nega-
tively supercoiled DNA.

19.3 DNA Can Be Supercoiled

A circular DNA molecule with the B conformation has an average of 10.4 base pairs per
turn. It is said to be relaxed if such a molecule would lie flat on a surface. This relaxed
double helix can be overwound or underwound if the strands of DNA are broken and
the two ends of the linear molecule are twisted in opposite directions. When the strands
are rejoined to create a circle, there are no longer 10.4 base pairs per turn as required to
maintain the stable B conformation. The circular molecule compensates for over- or
underwinding by forming supercoils that restore 10.4 base pairs per turn of the double
helix (Figure 19.19). A supercoiled DNA molecule would not lie flat on a surface. Each
supercoil compensates for one turn of the double helix.

)

Closed, circular DNA
with no supercoils

All base paired

Locally unwound region

DNA with two
negative supercoils
and n turns of the helix.

Closed, circular DNA with
no supercoils, n-2 turns of the
helix, and a locally unwound region.

19.4 Cells Contain Several Kinds of RNA 587

Most circular DNA molecules are supercoiled in cells but even long, linear DNA
molecules contain locally supercoiled regions. Bacterial chromosomes typically have
about five supercoils per 1000 base pairs of DNA. The DNA in the nuclei of eukaryotic
cells is also supercoiled as we will see in Section 19.5. All organisms have enzymes that
can break DNA, unwind or overwind the double helix, and rejoin the strands to alter
the topology. These enzymes, called topoisomerases, are responsible for adding and
removing supercoils. An example of a topoisomerase bound to DNA is shown in
Figure 19.20. These remarkable enzymes cleave one or both strands of DNA, unwind or
overwind DNA by rotating the cleaved ends, and then rejoin the ends to create (or remove)
supercoils.

One of the important consequences of supercoiling is shown in Figure 19.19. If
DNA is underwound, it compensates by forming negative supercoils in order to main-
tain the stable B conformation. (Overwinding produces positive supercoils.) An alterna-
tive conformation is shown on the right in Figure 19.19. In this form, most of the DNA
is double-stranded but there is a locally unwound region that is due to the slight under-
winding. The negatively supercoiled and locally unwound conformations are in equilib-
rium with the supercoiled form in excess because it is slightly more stable. The differ-
ence in free energy between the two conformations is quite small.

Most of the DNA in a cell is negatively supercoiled. This means that it is relatively
easy to unwind short regions of the molecule — especially those regions that are A/T-rich.
As mentioned earlier, localized unwinding is an essential step in the initiation of DNA
replication, recombination, repair, and transcription. Thus, negative supercoiling plays
an important biological role in these processes by storing the energy needed for local
unwinding. This is why topoisomerases that catalyze supercoiling are essential enzymes
in all cells.

19.4 Cells Contain Several Kinds of RNA

RNA molecules participate in several processes associated with gene expression. RNA
molecules are found in multiple copies and in several different forms within a given cell.
There are four major classes of RNA in all living cells:

1. Ribosomal RNA (rRNA) molecules are an integral part of ribosomes (intracellular
ribonucleoproteins that are the sites of protein synthesis). Ribosomal RNA is the
most abundant class of ribonucleic acid accounting for about 80% of the total
cellular RNA.

2. Transfer RNA (tRNA) molecules carry activated amino acids to the ribosomes for
incorporation into growing peptide chains during protein synthesis. tRNA mole-
cules are only 73 to 95 nucleotide residues long. They account for about 15% of the
total cellular RNA.

3. Messenger RNA (mRNA) molecules encode the sequences of amino acids in pro-
teins. They are the “messengers” that carry information from DNA to the transla-
tion complex where proteins are synthesized. In general, mRNA accounts for only
3% of the total cellular RNA. These molecules are the least stable of the cellular
ribonucleic acids.

4. Small RNA molecules are present in all cells. Some small RNA molecules have cat-
alytic activity or contribute to catalytic activity in association with proteins. Many
of these RNA molecules are associated with processing events that modify RNA
after it has been synthesized. Some are required for regulating gene expression.

RNAs are single-stranded molecules, but they often have complex secondary struc-
ture. Most single-stranded polynucleotides fold back on themselves to form stable
regions of base-paired, double-stranded RNA under physiological conditions. One type
of secondary structure is a stem-loop which forms when short regions of complementary
sequence form base pairs (Figure 19.21). The structure of the double-stranded regions
of such stem-loops resembles the structure of the A form of double-stranded DNA. As
we will see in Chapters 21 and 22, such structures are important in transcription and
are common features in transfer RNA, ribosomal RNA, and the small RNAs.

▲ Figure 19.20

Human ( Homo sapiens) topoisomerase I bound

to DNA. [PDB 1A31]

▲ Supercoiled telephone cords can be very
annoying.

KEY CONCEPT

Single-stranded RNA can fold back on
itself to create stable double-stranded
helical regions that resemble those in
DNA.

588 CHAPTER 19 Nucleic Acids

BOX 19.1 PULLING DNA

Single-molecule atomic-force spectroscopy is a powerful tool for
investigating the properties of single molecules. It has been used to
explore the properties of single-stranded DNA. The experiment in-
volves fixing one end of a single- stranded DNA molecule to a solid
surface and attaching the other end to a form of molecular tweezer
that can be used to pull the molecule and measure its resistance.

When this experiment is done with poly(dT) there is almost no
resistance until the molecule is in the fully extended form. This is
because poly(dT) has no significant secondary structure. However,
when poly(dA) is pulled there is initial resistance followed by a shift
to the fully extended form. Poly(dA) is helical in solution because the
adenylate residues stack on one another and the initial resistance is
due to breaking the helix.

The resistance can be measured and the calculated energy of
stacking is 15 kj mol -1 , in agreement with other determinations of
the stacking interactions of A bases on other As. The experiment
proves that stacking interactions are important in forming helical
DNA structures-even with single-stranded polynucleotides.

Pulling poly(dA). [Adapted from Ke et al. (2007)] ►

Pull

▲ Figure 19.21

Stem-loop structures in RNA. Single-stranded
polynucleotides, such as RNA, can form stem-
loops, or hairpins, when short regions of
complementary sequence form base pairs.
The stem of the structure consists of base-
paired nucleotides, and the loop consists of
noncomplementary nucleotides. Note that
the strands in the stem are antiparallel.

19.5 Nucleosomes and Chromatin

In 1879, ten years after Miescher’s discovery of nuclein, Walter Flemming observed
banded objects in the nuclei of stained eukaryotic cells. He called the material chromatin,
from the Greek chroma , meaning “color.” Chromatin is now known to consist of DNA
plus various proteins that package the DNA in a more compact form. Prokaryotic DNA
is also associated with protein to form condensed structures inside the cell. These struc-
tures differ from those observed in eukaryotes and are usually not called chromatin.

In a normal resting cell, chromatin exists as 30 nm fibers — long, slender threads
about 30 nm in diameter. In humans, the nucleus must accommodate 46 such chro-
matin fibers, or chromosomes. The largest human chromosome is about 2.4 x 10 8 bp; it
would be about 8 cm long if it were stretched out in the B conformation. During
metaphase (when chromosomes are most condensed) the largest chromosome is about
10 \x m long. The difference between the length of the metaphase chromosome and the
extended B form of DNA is 8000-fold. This value is referred to as the packing ratio.

A. Nucleosomes

The major proteins of chromatin are known as histones. Most eukaryotic species con-
tain five different histones — HI, H2A, H2B, H3, and H4. All five histones are small,
basic proteins containing numerous lysine and arginine residues whose positive charges
allow the proteins to bind to the negatively charged sugar-phosphate backbone of
DNA. The numbers of acidic and basic residues in typical mammalian histones are
noted in Table 19.4. Except for HI, the amino acid sequence of each type of histone is
highly conserved in all eukaryotes. For example, bovine histone H4 differs from pea
histone H4 in only two residues out of 102. Such similarity in primary structure implies
a corresponding conservation in tertiary structure and function.

Chromatin unfolds when it is treated with a solution of low ionic strength (<5 mM).
The extended chromatin fiber looks like beads on a string in an electron micrograph
(Figure 19.22). The “beads” are DNA-histone complexes called nucleosomes and the
“string” is double-stranded DNA.

19.5 Nucleosomes and Chromatin 589

Table 19.4 Basic and acidic residues in mammalian histones

Type

Molecular

weight

Number
of residues

Number
of basic
residues

Number
of acidic
residues

Rabbit thymus HI

21,000

213

Calf thymus H2A

14,000

129

Calf thymus H2B

1 3,800

125

Calf thymus H3

15,300

135

Calf thymus H4

11,300

102

Each nucleosome is composed of one molecule of histone HI, two molecules each
of histones H2A, H2B, H3, and H4, and about 200 bp of DNA (Figure 19.23). The H2A,
H2B, H3, and H4 molecules form a protein complex called the histone octamer around
which the DNA is wrapped. About 146 bp of DNA are in close contact with the histone
octamer forming a nucleosome core particle. The DNA between particles is called linker
DNA; it is about 54 bp long. Histone HI can bind to the linker DNA and to the core
particle but in the extended beads-on-a-string conformation HI is often absent. His-
tone HI is responsible for higher-order chromatin structures.

The structure of the nucleosome core particle has been determined by X-ray crys-
tallography (Figure 19.24). The eight histone subunits are arranged symmetrically as
four dimers: two H2A/H2B dimers and two H3/H4 dimers. The particle is shaped like a
flat disk with positively charged grooves that accommodate the sugar-phosphate back-
bone of DNA.

DNA wraps around the core particle forming about l 3 /4 turns per nucleosome. If
this DNA were in an extended conformation it would be about 50 nm in length but
when bound to the nucleosome core particle, the overall length is reduced to the width
of the disk, about 5 nm. The coils of DNA are topologically equivalent to negative su-
percoils and that’s why eukaryotic DNA becomes supercoiled when histones are re-
moved from chromatin.

The N- termini of all four core histones are rich in positively charged lysine (K) and
arginine (R) residues. These ends extend outward from the core particle where they inter-
act with DNA and negatively charged regions of other proteins (Figure 19.24). These in-
teractions serve to stabilize higher- order chromatin structures such as the 30 nm fiber.

▲ Figure 19.22

Electron micrograph of extended chromatin
showing the “beads-on-a-string” organization.

KEY CONCEPT

The vast majority of eukaryotic DNA is
bound to nucleosome core particles
spaced 200 bp apart.

▲ Figure 19.23

Diagram of nucleosome structure, (a) Histone octamer. (b) Nucleosomes. Each nucleosome is composed of a core particle plus histone HI and linker
DNA. The nucleosome core particle is composed of a histone octamer and about 146 bp of DNA. Linker DNA consists of about 54 bp. Histone HI
binds to the core particle and to linker DNA.

590 CHAPTER 19 Nucleic Acids

▲ Figure 19.24

Structure of the chicken {Gallus gallus) nucle-
osome core particle, (a) Histone octamer.

(b) Histone octamer bound to DNA — side
view showing the disk shape of the particle.
[PDB 1EQZ].

Specific lysine residues in these N-terminal ends can be acetylated by enzymes
known as histone acetyltransferases (HATS). For example, residues 5, 8, 12, 16, and 20
in histone H4 can be modified by acetylation.

SGRGKGGKGLGKGGAKRHRKVLR D.... (19.1)

5 8 12 16 20

Acetylation decreases the net positive charge of the histone N-termini and weakens the
interactions with other nucleosomes and proteins. The net result is a loosening up of
higher-order structures. Acetylation is associated with gene expression. HATS are pref-
erentially directed to sites where chromatin must be unraveled in order to transcribe a
gene. The relationship between transcriptional activation and histone acetylation is
under active investigation in many laboratories (Section 21.5C).

Histone deacetylases are responsible for removing acetyl groups from lysine
residues. This restores the positively charged side chains and allows nucleosomes
to adopt the more compact chromatin structure characteristic of regions that are not
expressed.

^ -ch 2 — ch 2 — ch 2 — ch 2 — nh 3

Acetylation

Deacetylation

' /vw 'CH 2 — CH 2 — CH 2 — CH 2 — NH — C — CH 3

(19.2)

B. Higher Levels of Chromatin Structure

The packaging of DNA into nucleosomes reduces the length of a DNA molecule
about tenfold. Further reduction comes from higher levels of DNA packaging. For
example, the beads-on-a-string structure is itself coiled into a solenoid to yield the
30 nm fiber. One possible model of the solenoid is shown in Figure 19.25. The 30 nm
fiber forms when every nucleosome contains a molecule of histone HI and adjacent
molecules of HI bind to each other cooperatively bringing the nucleosomes together
into a more compact and stable form of chromatin. Condensation of the beads-on-
a-string structure into a solenoid achieves a further fourfold reduction in chromo-
some length.

Finally, 30 nm fibers are themselves attached to an RNA-protein scaffold that holds
the fibers in large loops. There may be as many as 2000 such loops on a large chromo-
some. The RNA-protein scaffold of a chromosome can be seen under an electron
microscope when histones have been removed (Figure 19.26). The attachment of DNA
loops to the scaffold accounts for an additional 200-fold condensation in the length
of DNA.

The loops of DNA are attached to the scaffold at their base. Because the ends are
not free to rotate, the loops can be supercoiled. (Some of the supercoils can be seen in
Figure 19.26b, but most of the DNA is relaxed because one of the strands is broken
during treatment to remove histones.)

C. Bacterial DNA Packaging

Histones are found only in eukaryotes but prokaryotic DNA is also packaged with pro-
teins in a condensed form. Some of these proteins are referred to as histone like proteins
because they resemble eukaryotic histones. In most cases, there are no defined nucleo-
some-like particles in prokaryotes and much of the DNA is not associated with protein.
Bacterial DNA is attached to a scaffold in large loops of about 100 kb. This arrangement
converts the bacterial chromosome to a structure known as the nucleoid.

19.6 Nucleases and Hydrolysis of Nucleic Acids 591

19.6 Nucleases and Hydrolysis
of Nucleic Acids

Enzymes that catalyze the hydrolysis of phosphodiesters in nucleic acids are collectively
known as nucleases. There are a variety of different nucleases in all cells. Some of them
are required for the synthesis or repair of DNA as we will see in Chapter 20 and others
are needed for the production or degradation of cellular RNA (Chapter 21).

Some nucleases act on both RNA and DNA molecules but many act only on RNA
and others only on DNA. The specific nucleases are called ribonucleases (RNases) and
deoxyribonucleases (DNases). Nucleases can be further classified as exonucleases or en-
donucleases. Exonucleases catalyze the hydrolysis of phosphodiester linkages to release
nucleotide residues from only one end of a polynucleotide chain. The most common
exonucleases are the 3' — > 5' exonucleases but there are some 5' —> 3' exonucleases.
Endonucleases catalyze the hydrolysis of phosphodiester linkages at various sites within
a polynucleotide chain. Nucleases have a wide variety of specificities for nucleotide
sequences.

Nucleases can cleave either the 3' - or the 5 '-ester bond of a 3 '-5' phosphodiester
linkage. One type of hydrolysis yields a 5 '-monophosphate and a 3 '-hydroxyl group;
the other type yields a 3 '-monophosphate and a 5 '-hydroxyl group (see Figure 19.27).
A given nuclease can catalyze one reaction or the other but not both.

A. Alkaline Hydrolysis of RNA

The difference between ribose in RNA and 2'-deoxyribose in DNA may seem trivial but
it greatly affects the properties of the nucleic acids. The 2 '-hydroxyl group of ribose can
form hydrogen bonds in some RNA molecules and it participates in certain chemical
and enzyme-catalyzed reactions.

The effect of alkaline solutions on RNA and DNA illustrates the differences in
chemical reactivity that result from the presence or absence of the 2 '-hydroxyl group.
RNA treated with 0.1 M NaOH at room temperature is degraded to a mixture of
2'- and 3 '-nucleoside monophosphates within a few hours whereas DNA is stable
under the same conditions. Alkaline hydrolysis of RNA (Figure 19.28) requires a
2 '-hydroxyl group. In the first and second steps, hydroxide ions act only as catalysts

30 nm

▲ Figure 19.25

A model of the 30 nm chromatin fiber. In this
model the 30 nm fiber is shown as a solenoid,
or helix, formed by individual nucleosomes.
The nucleosomes associate through contacts
between adjacent histone HI molecules.

▲ Figure 19.26

Electron micrographs of a histone-depleted chromosome, (a) In this view, the entire protein scaffold is visible, (b) In this magnification of a portion of
(a), individual loops attached to the protein scaffold can be seen.

592 CHAPTER 19 Nucleic Acids

5' — » 3' exonuclease
G 0— P=0

O = P — 0°

3' — > 5' exonuclease

▲ Figure 19.27

Nuclease cleavage sites. Exonucleases act
on one free end of a polynucleotide and
cleave the next phosphodiester linkage.
Endonucleases cleave internal phosphodi-
ester linkages. Cleavage at bond A
generates a 5'-phosphate and a 3'-hydroxyl
terminus. Cleavage at bond B generates a
3'-phosphate and a 5'-hydroxyl terminus.
Both DNA (shown) and RNA are substrates
of nucleases.

5 'CH 2 n

◄ Figure 19.28

Alkaline hydrolysis of RNA. In Step 1, a hydroxide ion
abstracts the proton from the 2'-hydroxyl group of a
nucleotide residue. The resulting 2'-alkoxide is a nu-
cleophile that attacks the adjacent phosphorus atom,
displacing the 5'-oxygen atom and generating a 2', 3'-
cyclic nucleoside monophosphate. The cyclic interme-
diate is not stable in alkaline solution, however, and a
second hydroxide ion catalyzes its conversion to either
a 2'- or 3'-nucleoside monophosphate (Step 2).

B represents a purine or pyrimidine base.

( 2 )

2 , ,3'-Cyclic nucleoside monophosphate
+

O — P = O

2'-Nucleoside

monophosphate

y 0—? = 0

o©

3'-Nucleoside

monophosphate

since removing a proton from water (to form the 5' -hydroxyl group in the first step
or the 2'- or 3 '-hydroxyl group in the second) regenerates one hydroxide ion for each
hydroxide ion consumed. Note that a 2 ',3 '-cyclic nucleoside monophosphate inter-
mediate forms. The polyribonucleotide chain rapidly depolymerizes as each phospho-
diester linkage is cleaved. DNA is not hydrolyzed under alkaline conditions because it
lacks the 2 '-hydroxyl group needed to initiate intramolecular transesterification. The
greater chemical stability of DNA is an important factor in its role as the primary
genetic material.

B. Hydrolysis of RNA by Ribonuclease A

Bovine pancreatic ribonuclease A (RNase A) consists of a single polypeptide chain of
124 amino acid residues cross-linked by four disulfide bridges. (This is the same en-
zyme that we encountered in Chapter 4 in our discussion of disulfide bond formation

19.6 Nucleases and Hydrolysis of Nucleic Acids 593

and protein folding.) The enzyme has a pH optimum of about 6. RNase A catalyzes
cleavage of phosphodiester linkages in RNA molecules at 5 '-ester bonds. Cleavage occurs
to the right of pyrimidine nucleotide residues when chains are drawn in the 5 ' — > 3 '
direction. Thus, RNase A catalyzed hydrolysis of a strand with the sequence
pApG pUpApCpGpU yields pApGpUp + ApCp + GpU.

RNase A contains three ionic amino acid residues in the active site — Lys-41, His-
12, and His- 119 (Figure 19.29). Many studies have led to formulation of the mecha-
nism of catalysis shown in Figure 19.30. RNase A uses three fundamental catalytic
mechanisms: proximity (in the binding and positioning of a suitable phosphodiester
between the two histidine residues); acid-base catalysis (by His-119 and His-12); and
transition- state stabilization (by Lys-41). As in alkaline hydrolysis of RNA, hydrolysis
produces a leaving group with a 5 '-hydroxyl group and a 3 '-nucleoside monophos-
phate product. Water enters the active site on departure of the first product (P x ). Note
that in the RNase A-catalyzed reaction, the phosphate atom in the transition state is
pentacovalent. The pyrimidine binding pocket of the enzyme accounts for the speci-
ficity of RNase A.

Alkaline hydrolysis and the reaction catalyzed by RNase A differ in two important
ways. First, alkaline hydrolysis can occur at any residue whereas enzyme -catalyzed
cleavage occurs only at pyrimidine nucleotide residues. Second, hydrolysis of the cyclic
intermediate is random in alkaline hydrolysis (producing mixtures of 2'- and 3'-
nucleotides) but specific for RNase A-catalyzed cleavage (producing only 3 '-nucleotides).

C. Restriction Endonucleases

Restriction endonucleases are an important subclass of endonucleases that act on DNA.
The term restriction endonuclease is derived from the observation that certain bacteria
can block bacteriophage (virus) infections by specifically destroying the incoming bac-
teriophage DNA. Such bacteria restrict the expression of foreign DNA.

Many species of bacteria synthesize restriction endonucleases that bind to and
cleave foreign DNA. These endonucleases recognize specific DNA sequences and they
cut both strands of DNA at the binding site producing large fragments that are rapidly
degraded by exonucleases. The bacteriophage DNA is cleaved and degraded before the
genes can be expressed.

The host cell has to protect its own DNA from cleavage by restriction endonucle-
ases. This is accomplished by covalent modification of the bases that make up the po-
tential restriction endonuclease binding site. The most common covalent modification
is specific methylation of adenine or cytosine residues within the recognition sequence
(Section 18.7). The presence of methylated bases at the potential binding site inhibits
cleavage of the host DNA by the restriction endonuclease. Methylation is catalyzed by a
specific methylase that binds to the same sequence of DNA recognized by the restriction
endonuclease. Thus, cells that contain a restriction endonuclease also contain a methy-
lase with the same specificity.

Normally, all DNA of the host cell is specifically methylated and therefore protected
from cleavage. Any unmethylated DNA that enters the cell is cleaved by restriction en-
donucleases. Following DNA replication, each site in the host DNA is hemimethy-
lated — bases on only one strand are methylated. Hemimethylated sites are high affinity
substrates for the methylase but are not recognized by the restriction endonuclease.
Thus, hemimethylated sites are rapidly converted to fully methylated sites in the host
DNA (Figure 19.31).

Most restriction endonucleases (also called restriction enzymes) can be classified as
either type I or type II. Type I restriction endonucleases catalyze both the methylation
of host DNA and the cleavage of unmethylated DNA at a specific recognition sequence.
Type II restriction endonucleases are simpler in that they can only cleave double-
stranded DNA at or near an unmethylated recognition sequence — they do not possess a
methylase activity. Separate restriction methylases catalyze methylation of host DNA at
the same recognition sequences. The source of the methyl group in these reactions is
S - adenosylmethionine.

▲ Figure 19.29

The active site of bovine pancreatic RNase A.

(a) The active site of the enzyme has three
catalytic residues, His-12, His-119, and
Lys-41, whose side chains project into the
site where RNA will bind, (b) This figure
shows RNase A bound to an artificial
substrate (3’-phosphothymidine (3'-5’)-
pyrophosphate adenosine 3’-phosphate)
that mimics RNA. [PDB 1U1B]

594 CHAPTER 19 Nucleic Acids

Figure 19.30 ►

Mechanism of RNA cleavage by RNase A. In Step 1, His-12
abstracts a proton from the 2'-hydroxyl group of a pyrimi-
dine nucleotide residue. The resulting nucleophilic oxygen
atom attacks the adjacent phosphorus atom. His-119 (as an
imidazolium ion) donates a proton to the 5'-oxygen atom of
the next nucleotide residue to produce an alcohol leaving
group, Pi. Step 2 produces a 2',3'-cyclic nucleoside
monophosphate. Water enters the active site on departure
of Pi and in Step 3, His-119 (now in its basic form)
removes a proton from water. The resulting hydroxide ion
attacks the phosphorus atom to form a second transition
state. In Step 4, the imidazolium form of His-12 donates a
proton to the 2'-oxygen atom, producing P 2 . Py represents
a pyrimidine base.

19.6 Nucleases and Hydrolysis of Nucleic Acids 595

Table 19.5 Specificities of some common restriction endonucleases

Source

Enzyme 3

Recognition

sequence b

Acetobacter pasteurianus

Apa 1

GGGCCIC

Bacillus amyloliquefaciens H

BamHI

GIGATCC

Eschericia coli RY1 3

EcoRl

GIAA*TTC

Eschericia coli R245

EcoRII

ICC*TGG

Haemophilus aegyptius

Hae III

GGICC

Haemophilus influenzae R^

Hind\\\

ANAGCTT

Haemophilus parainfluenzae

HpaU

CICGG

Klebsiella pneumoniae

Kpn\

GGTACIC

Nocardia otitidis-caviarum

Not\

GCIGGCCGC

Providencia stuartii 1 64

Pst\

CTGCA1G

Serratia marcescens S b

Sma\

CCCfGGG

Xanthomonas badrii

Xba\

TICTAGA

Xanthomonas hold col a

Xho\

CITCGAG

a The names of restriction endonucleases are abbreviations of the names of the organisms that produce them.
Some abbreviated names are followed by a letter denoting the strain. Roman numerals indicate the order of dis-
covery of the enzyme in that strain.

Recognition sequences are written 5'to3'. Only one strand is represented. The arrows indicate cleavage sites.
Asterisks represent known positions where bases can be methylated.

(a) H 3 C

N N ' /w ' 3'

N N 5'

CH 3

5'^NNGAATTC
3'^NN C T T A AG

Replication

Following DNA
replication, the
GAATTC site
is hemimethylated.

h 3 c

5 , ' aa/ ' N N
3 , ' /w ' N N

G A A T T C N N^
C T T AAG N N^

Methylation

A methylase catalyzes
methylation of the
second adenine residue
in the recognition site.

n 3 c

5 , ' /w ' N N
3 , ' /w ' N N

CH 3

GAATT CNN ^ 3'
CTTAAGNN — 5'

Hundreds of type I and type II restriction endonucleases have been characterized.
The specificities of a few representative enzymes are listed in Table 19.5. In nearly all
cases, the recognition sites have a twofold axis of symmetry; that is, the 5' — > 3' se-
quence of residues is the same in both strands of the DNA molecule. Consequently, the
paired sequences “read” the same in either direction — such sequences are known as
palindromes. (Palindromes in English include BIB, DEED, RADAR, and even MADAM
EM ADAM, provided we ignore punctuation and spacing.)

EcoRl was one of the first restriction endonucleases to be discovered. It is present in
many strains of E. coli. As shown in Table 19.5 and Figure 19.31, EcoRl has a palin-
dromic recognition sequence of 6 bp (the 5' — » 3' sequence is GAATTC on each strand).
EcoRl is a homodimer. It possesses a twofold axis of symmetry like its substrate (see
next section). In E. coli , the companion methylase to EcoRl converts the second adenine
within the recognition sequence to X^-methyladenine. Any double-stranded DNA mol-
ecule with an unmethylated GAATTC sequence is a substrate for EcoRl. The endonucle-
ase catalyzes hydrolysis of the phosphodiesters that link G to A in each strand, thus
cleaving the DNA.

Some restriction endonucleases (including EcoRl, BaraHI, and Hmdlll) catalyze
staggered cleavage, producing DNA fragments with single- stranded extensions (Table 19.5
and Figure 19.31). These single- stranded regions are called sticky ends because they are
complementary and can thus re-form a double-stranded structure. Other enzymes,
such as Hae III and SmaR produce blunt ends with no single-stranded extensions.

(b)

N N 3'

N N ' /w ' 5'

5'^NNGAATTC
3'^NN C T T AAG

Restriction

The endonuclease
recognizes the GAATT C
sequence and cleaves
both strands of the
foreign DNA to
produce fragments
with staggered ends.

G 3' 5'

A A T T C

C T T AA

5' 3'G

v/w'
vTv/X/' ^ 7

▲ Figure 19.31

Methylation and restriction at the EcoRl site.

(a) Methylation of adenine residues at the
recognition site, (b) Cleavage of unmethy-
lated DNA to produce sticky ends.

D. EcoRl Binds Tightly to DNA

Restriction endonucleases must bind tightly to DNA in order to recognize a specific
sequence and cleave at a specific site. The structure of EcoRl bound to DNA has been
determined by X-ray crystallography. As shown in Figure 19.32, each half of the EcoRl
homodimer binds to one side of the DNA molecule so that the DNA molecule is almost
surrounded. The enzyme recognizes the specific nucleotide sequence by contacting base
pairs in the major groove. The minor groove (in the middle of the structure shown in
Figure 19.32) is exposed to the aqueous environment.

Several basic amino acid residues line the cleft that is formed by the two EcoRl
monomers. The side chains of these residues interact electrostatically with the

596 CHAPTER 19 Nucleic Acids

▲ Figure 19.32

EcoR\ bound to DNA. EcoR\ is composed of two
identical subunits (purple and blue). The en-
zyme is bound to a fragment of DNA with the
sequence CGC GAATTC GCG (recognition se-
quence underlined), (a) Side view, (b) Top view.

Figure 19.33 ►

Restriction map of bacteriophage A showing
the sites of cleavage by some restriction
enzymes. There is a single site for the en-
zyme Apa\, for example. Digestion of phage
A DNA with this enzyme yields two frag-
ments of 10.0 and 38.4 kb, as shown in the
first lane of Figure 19.34.

sugar-phosphate backbones of DNA. In addition, two arginine residues (Arg-145 and
Arg-200) and one glutamate residue (Glu-144) in each EcoRl monomer form hydrogen
bonds with base pairs in the recognition sequence thus ensuring specific binding. Other
nonspecific interactions with the backbones further stabilize the complex.

EcoRl is typical of proteins that recognize and bind to a specific DNA sequence. The
DNA retains its B conformation although in some cases the helix is slightly bent. Recog-
nition of a specific nucleotide sequence depends on interactions between the protein
and the functional groups on the bases that are exposed in the grooves. In contrast,
histones are examples of proteins that bind nonspecifically to nucleic acids. Binding of
such proteins depends largely on weak interactions between the protein and the
sugar-phosphate backbones and not on direct contact with the bases. All proteins that
bind to specific DNA sequences will also bind non- specifically to DNA with lower affinity
(Sections 21.3, 21.7A).

19.7 Uses of Restriction Endonucleases

Restriction endonucleases were discovered more than 40 years ago earning Nobel Prizes
in 1978 for Werner Arbor, Daniel Nathans, and Hamilton Smith “for the discovery
of restriction enzymes and their application to problems of molecular genetics.” The
first purified enzymes rapidly became important tools used to manipulate DNA in the
laboratory.

A. Restriction Maps

One of the first uses of restriction enzymes was in developing restriction maps of DNA,
that is, diagrams of DNA molecules that show specific sites of cleavage. Such maps are
useful for identifying fragments of DNA that contain specific genes.

An example of a restriction map of bacteriophage A DNA is shown in Figure 19.33.
The DNA of bacteriophage A is a linear, double- stranded molecule approximately
48,400 bp (48 kb) long. By treating this DNA with various restriction enzymes and
measuring the sizes of the resulting fragments, it is possible to develop a map of the
cleavage sites. An example of such restriction digests is shown in Figure 19.34. The
information from many restriction digests is combined to produce a complete and
accurate map.

B. DNA Fingerprints

The technology required for mapping restriction endonuclease cleavage sites was devel-
oped in the 1970s. It soon became apparent that the procedure could be used to identify
the sites of mutations, or variations, in the genome of a population. For example, dif-
ferent strains of bacteriophage A have slightly different restriction maps because their
DNA sequences are not identical. One strain may have the sequence GGGCCC near the
left-hand end of its DNA and it is cleaved by Apal, producing the two fragments shown
in Figure 19.34. Another strain may have the sequence GGACCC at the same site. Since
this sequence is not a cleavage site for Apal y the restriction map of this strain differs
from that shown in Figure 19.33.

Variations in DNA sequence can be used to identify individuals in a large heteroge-
neous population. In humans, for example, regions of the genome that are highly variable
give restriction fragments that are as unique as fingerprints. Such DNA fingerprints can
be used in paternity disputes or criminal investigations to identify or exonerate suspects.

An example of the use of DNA fingerprinting in a rape case is shown in Figure 19.35.
DNAs isolated from the victim, from the evidence (semen), and from two suspects are

Kpn\

Xba I Xho I

10.0 1

6.9 !

1 C

. 5.5

9.5

15.0

A DNA
48.4 kb

19.7 Uses of Restriction Endonucleases 597

▲ Figure 19.34

Digestion of bacteriophage A DNA by four restriction endonucleases. A solution of DNA is treated with
an enzyme and then electrophoresed on an agarose gel, which separates fragments according to
size. The smallest fragments move fastest and are found at the bottom of the gel. (A fragment of
1.5 kb is not visible in this figure.) The restriction enzyme for each digest is indicated at the top
of the lane. The lane at the right contains intact phage A DNA and a mixture of fragments from the
four digests. In the Xba\ digest, two fragments of 23.9 and 24.5 kb are not well resolved.

digested with a restriction endonuclease. The fragments are separated on an agarose gel
as described in Figure 19.34. This DNA is then transferred (blotted) to a membrane of
nylon. The bound DNA is denatured and exposed to small fragments of radioactively
labeled DNA from a variable region of the human genome. The labeled DNA probe hy-
bridizes specifically to the restriction fragments on the nylon membrane that are de-
rived from this region. The labeled fragments are identified by autoradiography.

The technique identifies suspect A as the rapist. In actual criminal investigations, a
number of different probes are used in combination with different restriction digests in
order to ensure that the pattern detected is unique. Modern techniques are powerful
and accurate enough to conclusively rule out some suspects and convict others. When
combined with polymerase chain reaction (PCR) amplification of DNA (Chapter 22), a
fingerprint can be obtained from a hair follicle or a tiny speck of blood.

C. Recombinant DNA

The discovery of restriction endonucleases soon led to the creation of recombinant DNA
molecules by joining, or recombining, different fragments of DNA produced by the en-
zymes. A common experiment involves excising a DNA fragment containing a target
gene of interest and inserting it into a cloning vector. Cloning vectors can be plasmids,
bacteriophage, viruses, or even small artificial chromosomes. Most vectors contain se-
quences that allow them to be replicated autonomously within a compatible host cell.

All cloning vectors have in common at least one unique cloning site, a sequence
that can be cut by a restriction endonuclease to allow site-specific insertion of foreign
DNA. The most useful vectors have several restriction sites grouped together in a multi-
ple cloning site called a polylinker.

► Stanley N. Cohen (1935-) (top) and Herbert Boyer (1936-) (bottom), who constructed the first
recombinant DNA using bacterial DNA and plasmids.

Victim

DNA size
markers

Blood

samples

Suspect
A

— I Suspect

Sexual

assault

evidence

r L

Female

fraction

Male

fraction

▲ Figure 19.35
DNA fingerprinting.

598 CHAPTER 19 Nucleic Acids

Figure 19.36 ►

Use of restriction enzymes to generate recom-
binant DNA. The vector DNA and the target
DNA are cleaved by restriction endonucle-
ases to generate ends that can be joined
together. In cases where sticky ends are pro-
duced, the two molecules join by annealing
(base pairing) of the complementary ends.
The molecules are then covalently attached
to one another in a reaction catalyzed by
DNA ligase.

Recombinant
DNA molecule

Fragments of DNA to be inserted into a vector can be generated by a variety of
means. For example, they can be produced by the mechanical shearing of long DNA
molecules or by digesting DNA with type II restriction endonucleases. Unlike shearing,
which cleaves DNA randomly, restriction enzymes cleave DNA at specific sequences.
For cloning purposes, this specificity offers extraordinary advantages.

The most useful restriction endonucleases produce fragments with single-stranded
extensions at their 3' or 5' ends. These sticky ends can transiently form base pairs to
complementary sticky ends on vector DNA and can be covalently joined to the vector in
a reaction catalyzed by DNA ligase (described in Section 20. 3C). Thus, the simplest
kinds of recombinant DNA are those constructed by digesting both the vector and the
target DNA with the same enzyme because the resulting fragments can be joined directly
by ligation (Figure 19.36).

Summary

1. Nucleic acids are polymers of nucleotides that are phosphate es-
ters of nucleosides. The amino and lactam tautomers of the bases
form hydrogen bonds in nucleic acids.

2. DNA contains two antiparallel strands of nucleotide residues
joined by 3 '-5' phosphodiester linkages. A and G in one strand
pair with T and C, respectively, in the other strand.

3. The double-helical structure of DNA is stabilized by hydrogen
bonding, hydrophobic effects, stacking interactions, and
charge-charge interactions. G/C-rich DNA is more difficult to de-
nature than A/T-rich DNA because the stacking interactions of
G/C base pairs are greater than those of A/T base pairs.

4. The most common conformation of DNA is called B-DNA; alter-
native conformations include A-DNA and Z-DNA.

5. Overwinding or underwinding the DNA helix can produce super-
coils that restore the B conformation. Negatively supercoiled
DNA exists in equilibrium with DNA that has locally unwound

6. The four major classes of RNA are ribosomal RNA, transfer RNA,
messenger RNA, and small RNA. RNA molecules are single-
stranded and have extensive secondary structure.

7. Eukaryotic DNA molecules are packaged with histones to form
nucleosomes. Further condensation and attachment to the scaf-
fold of a chromosome achieves an overall 8000-fold reduction in
the length of the DNA molecule in metaphase chromosomes.

8. The phosphodiester backbones of nucleic acids can be hydrolyzed
by the actions of nucleases. Alkaline hydrolysis and RNase A-
catalyzed hydrolysis of RNA proceed via a 2 ',3 '-cyclic nucleoside
monophosphate intermediate.

9. Restriction endonucleases catalyze hydrolysis of DNA at specific
palindromic nucleotide sequences. Specific methylases protect re-
striction sites from cleavage.

10. Restriction enzymes are useful for constructing restriction maps
of DNA, for DNA fingerprint analysis, and for constructing re-
combinant DNA molecules.

regions.

Selected Readings 599

Problems

1. Compare hydrogen bonding in the a helix of proteins to hydro-
gen bonding in the double helix of DNA. Include in the answer
the role of hydrogen bonding in stabilizing these two structures.

2. A stretch of double- stranded DNA contains 1000 bp and its base
composition is 58% (G + C). How many thymine residues are in
this region of DNA?

3. (a) Do the two complementary strands of a segment of DNA

have the same base composition?

(b) Does (A + G) equal (C + T)?

4. If one strand of DNA has the sequence

ATCGCGTAACATGGATTCGG

write the sequence of the complementary strand using the stan-
dard convention.

5. Poly A forms a single-stranded helix. What forces stabilize this
structure?

6. The imino tautomer of adenine occurs infrequently in DNA but
when it does it can pair with cytosine instead of thymine. Such
misp airing can lead to a mutation. Draw the adenine imino tau-
tomer/cytosine base pair.

7. Single- stranded poly-dA can hybridize to single- stranded poly-
dT to form Watson-Crick base-paired double-stranded DNA.
Under appropriate conditions a second strand of poly-dT can
bind in the major groove and form a triple-stranded DNA helix
with hydrogen bonds between the thymine and the N7 and amino
group in adenine. What would a plot of absorbance at 260 nm vs.
temperature look like for this unusual triple-stranded DNA?

8. Write the sequence of the RNA shown in Figure 19.21. Is it a
palindrome?

9. Consider a processive exonuclease that binds exclusively to double-
stranded DNA and degrades one strand in the 5' — » 3' direction.
In a reaction where the substrate is a 1 kb fragment of linear
DNA, what will be the predominant products after the digestion
has gone to completion?

10. The average molecular weight of a base pair in double-stranded
DNA is approximately 650 kDa. Using the data from Table 19.4,
calculate the mass ratio of protein to DNA in a typical 30 nm
chromatin fiber.

11. The human haploid genome contains 3.2 x 10 9 base pairs. How
many nucleosomes did you inherit from your mother?

12. A DNA molecule with the sequence pdApdGpdTpdC can be
cleaved by exonucleases. List the products of a single reaction cat-
alyzed by the following enzymes:

(a) a 3' — » 5' exonuclease that cleaves the 3' ester bond of a
phosphodiester linkage

(b) a 5' — » 3' exonuclease that cleaves the 5' ester bond of a
phosphodiester linkage

13. A non-sequence-specific endonuclease purified from Aspergillus
oryzae digests single-stranded DNA. Predict the effect of adding this
enzyme to a preparation of negatively supercoiled plasmid DNA.

14. One of the proteins in rattlesnake venom is an enzyme named
phosphodiesterase. Could polynucleotides be a substrate for this
enzyme? Why or why not?

15. RNase T1 cleaves RNA after G residues to leave a 3' phosphate
group. Predict the cleavage products of this substrate:

pppApCpUpCpApUpApGpCpUpApUpGpApGpU

16. How could bacteriophages escape the effects of bacterial restric-
tion endonucleases?

17. The free-living soil nematode C. elegans was the first metazoan to
have its entire 100 Mb genome sequenced. Overall, the worm
genome is 36% (G + C) and 64% (A + T). The restriction
endonuclease Hindlll recognizes and cuts the hexameric palin-
dromic sequence AAGCTT to generate sticky ends, (a) Approxi-
mately how many Hindlll sites would you expect to find in the
C. elegans genome? (b) If the worm genome was actually 25% G
and 25% A, approximately how many Hindlll sites would you
expect to find?

18. The recognition sites for the restriction endonucleases Bglll and
BamHl are shown below. Why is it possible to construct recombi-
nant DNA molecules by combining target DNA cut with Bglll
and a vector cut with BamHl?

l i

AGATCT GGATCC

Bglll BamHl

19. One of the E. coli host strains commonly used in recombinant
DNA technology carries defective genes for several restriction en-
donucleases. Why is such a strain useful?

Selected Readings

Historical Perspective

Clayton, J., and Denis. C. (eds.) (2003). 50 Years of DNA. (New York:
Nature/Pallgrave/Macmillan) .

fudson, H. F. (1996). The Eighth Day of Creation: Makers of the Revolution in
Biology , expanded ed. (Cold Spring Harbor, NY: Cold Spring Harbor
Laboratory Press).

Maddox, B. (2002). Rosalind Franklin: The Dark Lady of DNA
(New York: Perennial/HarperCollins).

Watson, J. D., and Berry, A. (2003). DNA: The Secret of Life (New York: Alfred
A. Knopf).

Watson, J. D. (1968). The Double Helix (New York: Atheneum).

Polynucleotide Structure and Properties

Berger, J. M., and Wang, J. C. (1996). Recent developments in DNA topoiso-
merase II structure and mechanism. Curr. Opin. Struct. Biol. 6:84-90.

Ferre-D’Amare, A. R., and Doudna, J. A. (1999). RNA FOLDS: insights from
recent crystal structures. Annu. Rev. Biophys. Biomol. Struct. 28:57-73.

Herbert, A., and Rich, A. (1996). The biology of left-handed Z-DNA. J. Biol.
Chem. 271:11595-11598.

Hunter, C. A. (1996). Sequence-dependent DNA structure. BioEssays 18:157-162.

Ke, C., Humeniuk, M., S-Gracz, H., and Marszalek, P. E. (2007). Direct meas-
urements of base stacking interactions in DNA by single-molecule atomic-
force spectroscopy. Phys. Rev. Lett. 99: 018302.

600 CHAPTER 19 Nucleic Acids

Kool, E. T., Morales, J. C., and Guckian, K. M. (2000). Mimicking the structure
and function of DNA: insights into DNA stability and replication. Angew.
Chem. Int. Ed. 39:990-1009.

Packer, M. J., and Hunter, C. A. (1998). Sequence- dependent DNA structure:
the role of the sugar-phosphate backbone. /. Mol. Biol. 280:407-420.

Saenger, W. (1984). Principles of Nucleic Acid Structure (New York:
Springer-Verlag).

Sharma, A., and Mondragon, A. (1995). DNA topoisomerases. Curr. Biol.
5:39-47.

Wang, J. C. (2009). A journey in the world of DNA rings and beyond. Annu.
Rev. Biochem. 78:31-54.

Chromatin

Bendich, A. J., and Drlica, K. (2000). Prokaryotic and eukaryotic chromosome:
what’s the difference? BioEssays 22:481-486.

Burlingame, R. W., Love, W. E., Wang, B.-C., Hamlin, R., Xuong, N.-H.,
and Moudrianakis, E. N. (1985). Crystallographic structure of the
octameric histone core of the nucleosome at a resolution of 3.3 A. Science
228:546-553.

Grigoryev, S. A., Arya, G., Correll, S., Woodcock, C. L., and Schlick, T. (2009).
Evidence for heteromorphic chromatin fibers from analysis of nucleosome in-
teractions. Proc. Natl. Acad. Sci. (USA) 106:13317-13322.

Kornberg, R. D. (1999). Twenty- five years of the nucleosome, fundamental
particle of the eukaryotic chromosome. Cell 98:285-294.

Ramakrishnan, V. (1997). Histone structure and the organization of the nucle-
osome. Annu. Rev. Biophys. Biomol. Struct. 26:83-112.

Richmond, T. J., Finch, J. T., Rushton, D., Rhodes, D., and Klug, A. (1984). Struc-
ture of the nucleosome core particle at 7 A resolution. Nature 31 1:532-537.

Van Holde, K., and Zlatanova, J. (1999). The nucleosome core particle: does it
have structural and functional relevance? BioEssays 21:776-780.

Workman, J. L., and Kingston, R. E. (1998). Alteration of nucleosome
structure as a mechanism of transcriptional regulation. Annu. Rev.
Biochem. 67:545-579.

Restriction Endonucleases

Kovall, R. A., and Mathews, B. W. (1999). Type II restriction endonucleases:
structural, functional and evolutionary relationships. Curr. Opin. Chem. Biol.
3:587-583.

McClarin, J. A., Frederick, C. A., Wang, B.-C., Greene, P., Boyer, H., Grable, J.,
and Rosenberg, J. M. (1986). Structure of the DNA-EcoRI endonuclease recog-
nition complex at 3 A resolution. Science 234:1526-1541.

Ne, M. (2000). Type I restriction systems: sophisticated molecular machines
(a legacy of Bertani and Weigle). Microbiol. Mol. Rev. 64:412-434.

o c

_ o

° o o o

° o

o o

° c

o o

DNA Replication, Repair,
and Recombination

T he transfer of genetic information from one generation to the next has puzzled
biologists since the time of Aristotle. Today, almost 2500 years later, we can ex-
plain why “like begets like ” Since genetic information is carried in DNA, it fol-
lows that the transfer of information from a parental cell to two daughter cells requires
exact duplication of DNA, a process known as DNA replication.

The DNA structure proposed by Watson and Crick in 1953 immediately suggested
a method of replication. The nucleotide sequence of one strand automatically specifies
the sequence of the other since the two strands of double-helical DNA are complemen-
tary. Watson and Crick proposed that the two strands of the helix unwind during DNA
replication and that each strand of DNA acts as a template for the synthesis of a com-
plementary strand. In this way, DNA replication produces two double- stranded daugh-
ter molecules, each containing one parental strand and one newly synthesized strand.
This mode of replication is termed semiconservative because one strand of the parental
DNA is conserved in each daughter molecule (Figure 20.1, on the next page).

In a series of elegant experiments, Matthew Meselson and Franklin W. Stahl
showed in 1958 that DNA was indeed replicated semiconservatively as predicted by
Watson and Crick. About the same time, reports of the purification and properties of
some of the enzymes involved in replication began to appear. The first DNA polymerase
was purified in 1958 by Arthur Kornberg, who was awarded the Nobel Prize for this
achievement. More recently, biochemists have isolated and characterized enzymes that
catalyze all the steps in DNA replication and have identified the genes that encode these
proteins. The actual mechanism of replication is much more complex — and more
interesting — than the simple scheme shown in Figure 20.1.

Establishing the steps of the replication mechanism required a combination of
both biochemical and genetic analysis. Much of what we know about DNA replication

The structure of DNA proposed by
Watson and Crick brought forth a
number of proposals as to how such
a molecule might replicate. These
proposals make specific predictions
concerning the distribution of parental
atoms among progeny molecules.
The results presented here give a de-
tailed answer to the question of this
distribution and simultaneously di-
rect our attention to other problems
whose solution must be the next step
in progress toward a complete un-
derstanding of the molecular basis of
DNA duplication.

— Matthew Meselson and
Franklin W. Stahl (1958)

Top: Holliday junction, an intermediate formed during recombination between two double-stranded DNA molecules.

601

602 CHAPTER 20 DNA Replication, Repair, and Recombination

Parental Daughter molecules

molecule , 1 ,

■ Parental strand

Newly synthesized strand

Origin

Termination site

Replisome

◄ Figure 20.1

Semiconservative DNA replication. Each strand of DNA acts as a template for synthesis of a new strand.
Each daughter molecule of DNA contains one parental strand and one newly synthesized strand.

has come from studies of the enzymes in Escherichia coli and its bacteriophages. The re-
sults of these studies have shown how large numbers of polypeptides assemble into
complexes that can carry out a complicated series of reactions. The DNA replication
complex is like a machine, or factory, whose parts are made of protein. Some of the
component polypeptides are partially active in isolation but others are active only in
association with the complete protein machine.

There are three distinct stages in DNA replication. (1) Initiation begins with the
correct assembly of the replication proteins at the site where DNA replication is to start.
(2) During the elongation stage, DNA is replicated semiconservatively as the complex
catalyzes the incorporation of nucleotides into the growing DNA strands. (3) Finally,
when replication terminates, the protein machine is disassembled and the daughter
molecules separate so that they can segregate into their new cells.

Protein machines that carry out a series of biochemical reactions are not confined
to the process of DNA replication but also occur in fatty acid synthesis (Section 16.1),
transcription (Chapter 21), and translation (Chapter 22). All four of these processes
include initiation, elongation, and termination steps. Furthermore, there is increasing
evidence that other processes of cellular metabolism are also carried out by complexes
of weakly associated enzymes and other macromolecules.

The maintenance of genetic information from generation to generation requires
that DNA replication be both rapid (because the entire complement of DNA must be
replicated before each cell division) and accurate. All cells have enzymes that correct
replication errors and repair damaged DNA. Furthermore, all cells can shuffle pieces of
DNA in a process known as genetic recombination. Both repair and recombination use
many of the same enzymes and proteins that are required for DNA replication.

The overall strategy of DNA replication, repair, and recombination in prokaryotes
and eukaryotes appears to be conserved, although specific enzymes vary among organ-
isms. Just as two different makes of automobile are similar even though individual parts
cannot be substituted for one another, so too are the mechanisms of DNA replication,
repair, and recombination similar in all organisms, even though the individual enzymes
may differ. We are going to focus on the biochemistry of these three processes in E. coli
because of its many well- characterized enzymes.

20.1 DNA Replication Is Bidirectional

The E. coli chromosome is a large, circular, double-stranded DNA molecule of
4.6 X 10 3 kilobase pairs (kb). Replication of this chromosome begins at a unique site
called the origin of replication and proceeds bidirectionally until the two replication
complexes meet at the termination site, where replication stops (Figure 20.2). The pro-
tein machine that carries out the polymerization reaction is called a replisome. It
contains a number of different proteins that are required for rapid and accurate DNA
replication. One replisome is located at each of the two replication forks, the points
where the parental DNA is unwound. Figure 20.3 shows an autoradiograph of a repli-
cating E. coli chromosome.

As parental DNA is unwound at a replication fork, each strand is used as a template
for the synthesis of a new strand. The rate of movement of a replication fork in E. coli is
approximately 1000 base pairs (bp) per second. In other words, each of the two new
strands is extended at the rate of 1000 nucleotides per second. Since there are two repli-
cation forks moving at this rate, the entire E. coli chromosome can be duplicated in
about 38 minutes.

◄ Figure 20.2

Bidirectional DNA replication in Escherichia coli. Semiconservative DNA replication begins at a
unique origin and proceeds in both directions. The synthesis of new strands of DNA (light gray)
occurs at the two replication forks where the replisomes are located. The two double-stranded DNA
molecules separate when the replication forks meet at the termination site. Note that each daugh-
ter molecule consists of one parental strand and one newly synthesized strand.

20.2 DNA Polymerase 603

Eukaryotic chromosomes are linear, double-stranded DNA
molecules that are usually much larger than the chromosomes of
bacteria. The large chromosomes of the fruit fly Drosophila
melanogaster for example, are about 5.0 X 10 4 kb in size or 10 times
larger than the E. coli chromosome. Replication in eukaryotes is also
bidirectional but whereas the E. coli chromosome has a unique origin
of replication, eukaryotic chromosomes have multiple sites where
DNA synthesis is initiated (Figure 20.4). The rate of fork movement
in eukaryotes is slower than in bacteria but the presence of many in-
dependent origins of replication enables the larger eukaryotic
genomes to be copied in approximately the same amount of time as
prokaryotic genomes.

20.2 DNA Polymerase

The synthesis of a new strand of DNA is achieved by the successive
addition of nucleotides to the end of a growing chain. This polymer-
ization is catalyzed by enzymes known as DNA- directed DNA poly-
merases, or simply DNA polymerases. E. coli cells contain three
different DNA polymerases; each protein is identified by a roman
numeral according to the order of its discovery. DNA polymerase I
repairs DNA and participates in the synthesis of one of the strands
of DNA during replication. DNA polymerase II plays a role in DNA
repair. DNA polymerase III is the major DNA replication enzyme
responsible for chain elongation during DNA replication and is the
essential part of the replisome.

DNA polymerase III contains ten different polypeptide subunits.

It is by far the largest of the three DNA polymerases (Table 20.1). The purified holoen-
zyme is an asymmetric dimer consisting of two copies of each polypeptide as shown in
Figure 20.5. The a , e, and 9 polypeptides combine to form two core complexes that are
responsible for the polymerization reactions. The /3 subunits form a sliding clamp that
surrounds each of the two DNA strands at the replication fork. Most of the remaining
subunits make up the y complex, or “clamp loader” that assists in assembly of the repli-
some and helps to keep the enzyme bound to parental DNA during successive polymer-
ization reactions.

▲ Figure 20.3

Autoradiograph of a replicating E. coli
chromosome. The DNA was labeled with
3 H-deoxythymidine, and the radioactivity
detected by overlaying the replicating chro-
mosome with photographic emulsion. The
autoradiograph shows that the E. coli chro-
mosome has two replication forks.

▲ Figure 20.4

Electron micrograph of replicating DNA from an embryo of the fruit fly Drosophila melanogaster. Note
the large number of replication forks at opposite ends of “bubbles” of duplicated DNA.

▲ Arthur Kornberg (1918-2007). Kornberg
received the Nobel Prize in 1959 for his
discovery of DNA polymerase.

604 CHAPTER 20 DNA Replication, Repair, and Recombination

KEY CONCEPT

Two replication forks move in
opposite directions from the origin
of replication.

The convention for assigning the direc-
tion of DNA strands is described in
Section 19.2A.

Figure 20.5 ►

Diagram of the subunit composition of E. coli
DNA polymerase III. The holoenzyme consists
of two core complexes (containing a, s and
6), paired copies of p and r, and a single y
complex (y, 8, 8' , with two copies each of ifj,
and %)■ The structure is thus an asymmetric
dimer. Other models of the holoenzyme
structure have been proposed. [Adapted
from O’Donnell, M. (1992). Accessory pro-
tein function in the DNA polymerase III
holoenzyme from E. coli. BioEssays
14:105-111.]

Table 20.1 Subunits of DNA polymerase III holoenzyme

Subunit

M r

Gene

Activity

1 30 000

polC/dnaE

Polymerase

27 000

? core

dnaQ/mutD

3' — » 5' exonuclease

8846 J

holE

40 000

dnaN

Forms sliding clamp

71 000

dnaX

Enhances dimerization of core; ATPase

47 000 >

| dnaX \

38 700

holA

8 '

36 900

/ 7

f complex

holS >

Enhance processivity; assist

16 600

holC

in replisome assembly

15 174 J

’ holD )

A. Chain Elongation Is a Nucleotidyl Group Transfer Reaction

All DNA polymerases, including DNA polymerase III, synthesize DNA by adding one
nucleotide at a time to the 3' end of the growing chain. The nucleotide substrate is a de-
oxyribonucleoside 5 '-triphosphate (dNTP). The specific nucleotide is determined by
Watson- Crick base pairing to the template strand; adenine (A) pairs with thymine (T)
and guanine (G) pairs with cytosine (C). Since the pool of each dNTP in a cell is
approximately equal, this means that on average the enzyme spends three quarters of its
time discriminating against incorrect dNTPs that have diffused into the catalytic site
where they try to base pair with the template strand.

DNA polymerase III catalyzes the formation of a phosphodiester linkage between
the incoming dNTP and the growing chain. The incoming dNTP forms a base pair with
a residue of the template strand (Figure 20.6). Once a correct base pair has formed, the
free 3 '-hydroxyl group of the nascent DNA chain carries out a nucleophilic attack on
the a-phosphorus atom of the incoming dNTP. This reaction leads to the addition of a
nucleoside monophosphate and displacement of pyrophosphate. Subsequent hydrolysis
of the pyrophosphate by the abundant enzyme pyrophosphatase makes the polymeriza-
tion reaction highly favorable in the direction of polymerization. The direction of poly-
merization (chain growth) is defined as 5' — » 3', reading across the carbon atoms on
the sugar ring of the newly added residue.

Core complex

20.2 DNA Polymerase 605

New strand
5' DNA

©■

o— P — 0— P — 0— P — O

©r

3' DNA
H O

V H H\

Hydrogen bonds

k tr\ -frv v m r\ r\

^ N ^ have formed

2 Pi «

Pyrophosphatase

h 2 o

ppi

Chain growth

5' — > 3'

◄ Figure 20.6

Elongation of a DNA chain. A base
pair is created when an incoming
deoxynucleoside 5'-triphosphate
(blue) forms hydrogen bonds with a
residue of the parental strand. A
phosphodiester linkage forms when
the terminal 3'-hydroxyl group
attacks the a-phosphorus atom of
the incoming nucleotide. Hydrolysis
of the released pyrophosphate
makes the overall reaction thermo-
dynamically favorable.

5' DNA

Template strand

OH H

606

CHAPTER 20 DNA Replication, Repair, and Recombination

DNA polymerase III advances by one residue, after each addition reaction, and an-
other nucleotidyl group transfer reaction occurs. This mechanism ensures that the new
chain is extended by the stepwise addition of single nucleotides that are properly
aligned by base pairing with the template strand. As expected, DNA polymerase III can-
not synthesize DNA in the absence of a template, nor can it add nucleotides in the ab-
sence of a 3' end of a preexisting chain. In other words, DNA polymerase III requires
both a template and a primer as substrates for synthesis to occur.

As noted earlier, DNA replication inside the cell proceeds at a rate of approximately
1000 nucleotides per second. This is the fastest known rate of any in vivo polymeriza-
tion reaction. The rate of polymerization catalyzed by purified DNA polymerase III in
vitro , however, is much slower, indicating that the isolated enzyme lacks some compo-
nents necessary for full activity. Only when the complete replisome is assembled does
polymerization in vitro occur at approximately the rate found inside the cell.

The elongation reaction in fatty acid syn-
thesis is another example of a proces-
sive polymerization reaction catalyzed
by a large complex (Section 16.1C).

The glycogen synthase reaction is an
example of a distributive polymerization
reaction (Section 12.5A).

B. DNA Polymerase III Remains Bound to the Replication Fork

Once DNA synthesis has been initiated, the polymerase remains bound at the replica-
tion fork until replication is complete. The 3' end of the growing chain remains associ-
ated with the active site of the enzyme while many nucleotides are added sequentially.
As part of the replisome, the DNA polymerase III holoenzyme is highly processive (see
Section 12. 5 A). This means that only a small number of DNA polymerase III molecules
are needed to replicate the entire chromosome. Processivity also accounts for the rapid
rate of DNA replication.

The processivity of the DNA polymerase III holoenzyme is due, in part, to the /3
subunits of the enzyme. These subunits have no activity on their own but when assem-
bled into the holoenzyme they form a ring that can completely surround the DNA mol-
ecule. The ring is formed by two /3 subunits that form a head-to-tail dimer. Each of the
subunits contains three similar domains consisting of a /3 sandwich fold with two a
helices at the interior edge that interact with DNA (Figure 20.7). The /3 subunits thus
act as a sliding clamp locking the polymerase onto the DNA substrate. Incorporating
DNA polymerase III into an even larger protein machine at the replication fork further
ensures that the enzyme remains associated with the nascent DNA chains during poly-
merization. Many other biochemically characterized DNA replication systems have
evolved the same strategy to make DNA replication faster (more efficient). For example,
two related bacteriophage, T 4 and RB69, both encode a replication accessory protein,
gp45, that forms a circular clamp (Figure 20.7). This clamp structure locks the phage-
encoded DNA polymerases onto their DNA substrates and enhances processivity.
Figure 20.8 shows a model for how this is likely to work in vivo for bacteriophage DNA
polymerase bound to DNA. The sliding clamp surrounds the double-stranded region of
DNA and interacts with the subunits containing the polymerase activity that bind to the
single-stranded region of the replication fork. Eukaryotic DNA polymerases use the
same strategy to clamp onto their substrates (see Section 20.6).

Figure 20.7 ►

DNA polymerases can use sliding ring clamps
to increase processivity. These three crystal
structures show the convergent evolution
of structure and function; (a) the p subunit
of E. coli DNA polymerase III [PDB 1MMI];
(b) Proliferating Cell Nuclear Antigen
(PCNA) that performs the same function in
archaebacteria; [PDB 3LX1] (c) gp45 from
bacteriophage T4 is also a sliding ring that
clamps DNA polymerase to its DNA sub-
strate. [PDB 1CZD]

20.3 DNA Polymerase Synthesizes Two Strands Simultaneously 607

RB69 DNA polymerase (gp43)

RB69 sliding clamp (gp45)

◄ Figure 20.8

Model of bacteriophage DNA polymerase
bound to DNA. The sliding clamp (purple)
surrounds the newly synthesized double-
stranded DNA. The subunit containing the
active site is shown in blue. The 3' end of
the nascent strand is positioned at the ac-
tive site and the single-stranded region of
the template strand extends leftward. The
DNA polymerase will move from right to
left as the nascent strand is extended.
[PDB 1WAI].

C. Proofreading Corrects Polymerization Errors

The DNA polymerase III holoenzyme also possesses a3'^5' exonuclease activity.
This exonuclease, whose active site lies primarily within the s subunit, can catalyze hy-
drolysis of the phosphodiester linkage that joins the 3 '-terminal residue to the rest of the
growing polynucleotide chain. Thus, the DNA polymerase III holoenzyme can catalyze
both chain elongation and degradation. The exonuclease activity allows the holoen-
zyme to proofread, or edit, newly synthesized DNA in order to correct any mismatched
base pairs. When DNA polymerase III recognizes a distortion in the DNA produced by
an incorrectly paired base, the exonuclease activity of the enzyme catalyzes removal of
the mispaired nucleotide before polymerization continues.

An incorrect base is incorporated about once every 10 5 polymerization steps
for an error rate of about 10 -5 . The 3' — > 5' proofreading exonuclease activity will re-
move 99% of these incorrect nucleotides. It has an error rate of 10 -2 . The combina-
tion of these two sequential reactions yields an error rate for polymerization of 10 -7 .
This is one of the lowest error rates of any enzyme. Most of these replication errors
are subsequently repaired by separate DNA repair enzymes (Section 20.7) yielding an
overall error rate for DNA replication of between 10 -9 and 10 -10 . Despite this im-
pressive accuracy, replication errors are common when large genomes are duplicated.
(Recall that the human genome contains 3.2 X 10 9 bp, which means that, on average,
each time the genome is replicated an error gets transmitted to one of the two daugh-
ter cells.) Mistakes that occur during DNA replication are the most common source
of mutation. What this means is that most of evolution is due to the inaccuracy of
DNA replication!

Proofreading is possible because the
polymerization mechanism is head
growth not tail growth (Box 12.3).

KEY CONCEPT

The accuracy of DNA polymerase
combined with proofreading and DNA
repair makes DNA replication the most
accurate biochemical reaction known.

20.3 DNA Polymerase Synthesizes Two
Strands Simultaneously

DNA polymerases catalyze chain elongation exclusively in the 5' — > 3' direction, as
shown in Figure 20.6. Since the two strands of DNA are antiparallel, 5' — > 3' synthesis
using one template strand occurs in the same direction as fork movement but 5' — > 3'
synthesis using the other template strand occurs in the direction opposite fork move-
ment (Figure 20.9). The new strand formed by polymerization in the same direction as

608 CHAPTER 20 DNA Replication, Repair, and Recombination

▲ Figure 20.9

Diagram of the replication fork. The two newly
synthesized strands have opposite polarity.
On the leading strand, 5' —> 3 ' synthesis
moves in the same direction as the replica-
tion fork; on the lagging strand, 5' — >3'
synthesis moves in the opposite direction.

| Parental DNA (unlabeled)

i— I Newly synthesized DNA
without 3 H label

|— | Newly synthesized DNA labeled
with 3 H-deoxythymidine

Separate by size (only newly
synthesized DNA containing
3 H-deoxythymidine is shown)

Long fragments from
leading strand

fork movement is called the leading strand. The new strand formed by polymerization
in the opposite direction is called the lagging strand. Recall that the DNA polymerase III
holoenzyme dimer contains two core complexes that can catalyze polymerization. One
of these is responsible for synthesis of the leading strand and the other is responsible for
synthesis of the lagging strand.

A. Lagging Strand Synthesis Is Discontinuous

The leading strand is synthesized as one continuous polynucleotide beginning at the
origin and ending at the termination site. In contrast, the lagging strand is synthesized
discontinuously in short pieces in the direction opposite fork movement. These pieces
of lagging strand are then joined by a separate reaction. In Section 20.4, we present a
model of the replication fork that explains how one enzyme complex can synthesize
both strands simultaneously.

An experiment that illustrates discontinuous DNA synthesis is shown in
Figure 20.10. E. coli DNA is labeled with a short pulse of 3 H-deoxythymidine. The
newly made DNA molecules are then isolated, denatured, and separated by size. The ex-
periment detects two types of labeled DNA molecules: very large DNA molecules that
collectively contain about half the radioactivity of the partially replicated DNA and
shorter DNA fragments of about 1000 residues that collectively contain the other half of
the radioactivity. The large DNA molecules arise from continuous synthesis of the lead-
ing strand while the shorter fragments arise from discontinuous synthesis of the lagging
strand. The short pieces of lagging strand DNA are named Okazaki fragments in honor of
their discoverer, Reiji Okazaki. The overall mechanism of DNA replication is called
semidiscontinuous to emphasize the different mechanisms for replicating each strand.

B. Each Okazaki Fragment Begins with an RNA Primer

It was clear that lagging strand synthesis is discontinuous but it was not obvious how
synthesis of each Okazaki fragment is initiated. The problem is that no DNA poly-
merase can begin polymerization de novo ; they can only add nucleotides to existing
polymers. This limitation presents little difficulty for leading strand synthesis since once
DNA synthesis is under way nucleotides are continuously added to a growing chain.
However, on the lagging strand the synthesis of each Okazaki fragment requires a new
initiation event. This is accomplished by making short pieces of RNA at the replication
fork. These RNA primers are complementary to the lagging strand template. Each
primer is extended from its 3' end by DNA polymerase to form an Okazaki fragment as
shown in Figure 20.11. (Synthesis of the leading strand also begins with an RNA primer
but only one primer is required to initiate synthesis of the entire strand.)

The use of short RNA primers gets around the limitation imposed by the mecha-
nism of DNA polymerase — namely, that it cannot initiate DNA synthesis de novo. The
primers are synthesized by a DNA-dependent RNA polymerase enzyme called
primase — the product of the dnaG gene in E. coli. The three-dimensional crystal struc-
ture of the DnaG catalytic domain revealed that its folding and active site are distinct
from the well studied polymerases suggesting that it may employ a novel enzyme mech-
anism. Primase is part of a larger complex called the primosome that contains many
other polypeptides in addition to primase. The primosome, along with DNA poly-
merase III, is part of the replisome.

As the replication fork progresses, the parental DNA is unwound and single-
stranded DNA becomes exposed. Primase catalyzes the synthesis of a short RNA primer
about once every second using this single-stranded DNA as a template. The primers are
only a few nucleotides in length. Since the replication fork advances at a rate of about

◄ Figure 20.10

Discontinuous DNA synthesis demonstrated by analysis of newly synthesized DNA. Nascent DNA mole-
cules are labeled in E. coli with a short pulse of 3 H-deoxythymidine. The cells are lysed, the DNA
is isolated, and single strands are separated by size. The labeled DNA molecules fall into two classes:
long molecules arising from continuous synthesis of the leading strand and short fragments arising
from discontinuous synthesis of the lagging strand.

20.3 DNA Polymerase Synthesizes Two Strands Simultaneously 609

1000 nucleotides per second, one primer is synthesized for approximately every 1000
nucleotides that are incorporated. DNA polymerase III catalyzes synthesis of DNA in
the 5' — > 3' direction by extending each short RNA primer.

C. Okazaki Fragments Are Joined by the Action of DNA
Polymerase I and DNA Ligase

Okazaki fragments are eventually joined to produce a continuous strand of DNA. The
reaction proceeds in three steps: removal of the RNA primer, synthesis of replacement
DNA, and sealing of the adjacent DNA fragments. The steps are carried out by the com-
bined action of DNA polymerase I and DNA ligase.

DNA polymerase I of E. coli was the enzyme discovered by Arthur Kornberg. It was
the first enzyme to be found that could catalyze DNA synthesis using a template strand.
In a single polypeptide, DNA polymerase I contains the two activities found in the DNA
polymerase III holoenzyme: 5' — > 3' polymerase activity and 3' — > 5' proofreading ex-
onuclease activity. In addition, DNA polymerase I has 5' — > 3' exonuclease activity, an
activity not found in DNA polymerase III.

DNA polymerase I can be cleaved with certain proteolytic enzymes to generate a
small fragment that contains the 5' — > 3' exonuclease activity and a larger fragment
that retains the polymerization and proofreading activities. The larger fragment con-
sists of the C-terminal 605 amino acid residues, and the smaller fragment contains the
remaining N-terminal 323 residues. The large fragment, known as the Klenow frag-
ment, was widely used for DNA sequencing and is still used in many other techniques
that require DNA synthesis without 5' — > 3' degradation. In addition, many studies of
the mechanisms of DNA synthesis and proofreading use the Klenow fragment as a
model for more complicated DNA polymerases.

Figure 20.12 shows the structure of the Klenow fragment complexed with a frag-
ment of DNA containing a mismatched terminal base pair. The 3' end of the nascent
strand is positioned at the 3' —> 5' exonuclease site of the enzyme. During polymerization,
the template strand occupies the groove at the top of the structure and at least 10 bp of
double- stranded DNA are bound by the enzyme as shown in the figure. Many of the
amino acid residues involved in binding DNA are similar in all DNA polymerases

Figure 20.12 ►

Structure of the Klenow fragment with a bound DNA fragment. The enzyme wraps around the DNA. The

3' end of the nascent strand is positioned at the 3' — » 5' exonuclease site (lower left). During DNA
synthesis in vivo the template strand extends beyond the double-stranded region shown in the crystal
structure. [PDB 1KLN].

◄ Figure 20.11

Diagram of lagging strand synthesis. A short
piece of RNA (brown) serves as a primer for
the synthesis of each Okazaki fragment. The
length of the Okazaki fragment is deter-
mined by the distance between successive
RNA primers.

3' -» 5' exonuclease
active site

610 CHAPTER 20 DNA Replication, Repair, and Recombination

▲ E. coli DNA ligase bound to nicked DNA.
[PDB 20W0]

▲ Structure of nicked DNA substrate when
bound by DNA ligase [PDB 20W0].

although the enzymes may be otherwise quite different in three-dimensional structure
and amino acid sequence.

The unique 5' — » 3' exonuclease activity of DNA polymerase I removes the RNA
primer at the beginning of each Okazaki fragment. (Since it is not part of the Klenow
fragment, the 5' — > 3' exonuclease is not shown in Figure 20.12, but it would be located
at the top of the structure next to the groove that accommodates the template strand.)
As the primer is removed, the polymerase synthesizes DNA to fill in the region between
Okazaki fragments, a process called nick translation (Figure 20.13). In nick translation,
DNA polymerase I recognizes and binds to the nick between the 3' end of an Okazaki
fragment and the 5' end of the next primer. The 5' — > 3' exonuclease then catalyzes hy-
drolytic removal of the first RNA nucleotide while the 5' — » 3' polymerase adds a de-
oxynucleotide to the 3' end of the DNA chain. In this way, the enzyme moves the nick
along the lagging strand. DNA polymerase I dissociates from the DNA after completing
10 or 12 cycles of hydrolysis and polymerization, leaving behind two Okazaki fragments
that are separated by a nick in the phosphodiester backbone. The removal of RNA
primers by DNA polymerase I is an essential part of DNA replication because the final
product must consist entirely of double-stranded DNA.

The last step in the synthesis of the lagging strand of DNA is the formation of a
phosphodiester linkage between the 3 '-hydroxyl group at the end of one Okazaki frag-
ment and the 5 '-phosphate group of an adjacent Okazaki fragment. This step is
catalyzed by DNA ligase. The DNA ligases in eukaryotic cells and in bacteriophage-
infected cells require ATP as a cosubstrate. In contrast, E. coli DNA ligase uses NAD® as
a cosubstrate. NAD® is the source of the nucleotidyl group that is transferred, first to
the enzyme and then to the DNA, to create an ADP-DNA intermediate. The proposed
mechanism of DNA ligase in E. coli is shown in Figure 20.14. The net reaction is

DNA (nicked) + NAD® > DNA(sealed) + NMN® + AMP (20.1)

20.4 Model of the Replisome

The replisome contains a primosome, the DNA polymerase III holoenzyme, and add-
itional proteins that are required for DNA replication. The assembly of many proteins
into a single machine allows coordinated synthesis of the leading and lagging strands at
the replication fork.

The template for DNA polymerase III is single-stranded DNA. This means that the
two strands of the parental double helix must be unwound and separated during repli-
cation. This unwinding is accomplished primarily by a class of proteins called helicases.
The helicase DnaB is required for DNA replication in E. coli. DnaB is one of the sub-
units of the primosome that, in turn, is part of the larger replisome. The rate of DNA
unwinding is directly coupled to the rate of polymerization as the replisome moves
along the chromosome. Unwinding is assisted by the actions of various topoisomerases
(Section 19.3) that relieve supercoiling ahead of and behind the replication fork. These
enzymes are not part of the replisome but they are required for replication. The most
important topoisomerase in E. coli is topoisomerase II, or gyrase. Mutants lacking this
enzyme cannot replicate their DNA. The end result is the production of two daughter
molecules each containing one newly synthesized stand and one parental strand as
shown in Figure 20.1. At no time during DNA replication is there a significant stretch of
single-stranded DNA other than that found on the lagging strand template.

Another protein that is part of the replisome is single-strand binding protein
(SSB), also known as helix-destabilizing protein. SSB binds to single-stranded DNA and
prevents it from folding back on itself to form double-stranded regions. SSB is a
tetramer of four identical small subunits. Each tetramer covers about 32 nucleotides of
DNA. Binding of SSB to DNA is cooperative; that is, binding of the first tetramer facili-
tates binding of the second, and so on. The presence of several adjacent SSB molecules
on single-stranded DNA produces an extended, relatively inflexible, DNA conformation.
Single-stranded DNA coated with SSB is an ideal template for synthesis of the comple-
mentary strand during DNA replication because it is free of secondary structure.

20.4 Model of the Replisome 611

(a) Completion of Okazaki fragment synthesis leaves a nick between the Okazaki < Figure 20.13

fragment and the preceding RNA primer on the lagging strand. Joining of Okazaki fragments by the combined

action of DNA polymerase I and DNA ligase.

Okazaki fragment

RNA primer

(b) DNA polymerase I extends the Okazaki fragment while its 5'— » 3' exonuclease
activity removes the RNA primer. This process, called nick translation, results in
movement of the nick along the lagging strand.

5'-> 3' polymerization

Nick DNA ligase

(d) DNA ligase catalyzes formation of a phosphodiester linkage, which seals the
nick, creating a continuous lagging strand. The enzyme then dissociates from
the DNA.

Closed nick

3 '

612 CHAPTER 20 DNA Replication, Repair, and Recombination

Sealed DNA strand

▲ Figure 20.14

Proposed mechanism of DNA ligase in E. coli. Using NAD© as a cosubstrate, E. coli DNA ligase catalyzes the formation of a phosphodiester linkage at
a nick in DNA. In Step 1, the e-amino group of a lysine residue of DNA ligase attacks the phosphorus atom bonded to the 5'-oxygen atom of the adeno-
sine moiety of NAD®. Nicotinamide mononucleotide (NMN©) is displaced, generating an AMP-DNA-ligase intermediate. (With DNA ligases that use
ATP as the cosubstrate, pyrophosphate is displaced.) In Step 2, an oxygen atom of the free 5'-phosphate group of the DNA attacks the phosphate
group of the AMP-enzyme complex, forming an ADP-DNA intermediate. In Step 3, the nucleophilic 3'-hydroxyl group on the terminal residue of the
adjacent DNA strand attacks the activated 5'-phosphate group of ADP-DNA, releasing AMP and generating a phosphodiester linkage that seals the
nick in the DNA strand. B represents any base.

A model of DNA synthesis by the replisome is shown in Figure 20.15. The primo-
some containing the primase and helicase is located at the head of the replication fork,
followed by a DNA polymerase III holoenzyme. (In order to simplify the figure, only
the core complexes of DNA polymerase III are shown.) Primase synthesizes an RNA
primer approximately once every second as the helicase unwinds the DNA. One of the
two core complexes in the holoenzyme dimer synthesizes the leading strand continuously

20.4 Model of the Replisome 613

▲ Model for E. coli SSB tetramer bound to
ssDNA [PDB 1EYG]

◄ DNA bound to SSB Model for the ex-
tended conformation of three SSB tetramers
bound cooperatively to ssDNA. [PDB 1EYG]

Source: Nature Structural and Molecular Biology
7:648-652 (2000) Raghunathan et al.

(a) The lagging-strand template loops back through the replisome so that the
leading and lagging strands are synthesized in the same direction. SSB binds
to single-stranded DNA.

RNA

primer

Primosome

/3-clamp

merase

complexes

SSB tetramer

(b) As helicase unwinds the DNA template, primase synthesizes an RNA primer.
The lagging-strand polymerase completes an Okazaki fragment.

5 '

◄ Figure 20.15

Simultaneous synthesis of leading and lagging
strands at a replication fork. The replisome
contains the DNA polymerase III holoen-
zyme (only the core complexes are shown);
a primosome containing primase, a helicase,
and other subunits; and additional compo-
nents including single-strand binding protein
(SSB). One core complex of the holoenzyme
synthesizes the leading strand while the
other core complex synthesizes the lagging
strand. The lagging-strand template
is looped back through the replisome so
that the leading and lagging strands can be
synthesized in the same direction as fork
movement, (c) and (d) continue on the next
page.

614

CHAPTER 20 DNA Replication, Repair, and Recombination

Figure 20.15 (Continued) ►

(d) The lagging-strand polymerase binds to a newly synthesized primer and begins
synthesizing another Okazaki fragment.

in the 5' —> 3' direction while the other extends the RNA primers to form Okazaki
fragments. The lagging-strand template is thought to fold back into a large loop. This
configuration allows both the leading and lagging strands to be synthesized in the same
direction as fork movement.

The two core complexes of the DNA polymerase III holoenzyme are drawn in the
model as equivalent but their roles in DNA replication are not equivalent. One of them
remains firmly bound to the leading-strand template whereas the other binds the
lagging-strand template until it encounters the RNA primer of the previously synthe-
sized Okazaki fragment. At this point the core complex releases the lagging-strand
template. The lagging-strand template reassociates with the holoenzyme at the site of
the next primer and synthesis continues (Figure 20.15d). The entire holoenzyme is
extremely processive since half of it remains associated with the leading strand from the
beginning of replication until termination while the other half processively synthesizes
stretches of 1000 nucleotides in the lagging strand. The y complex of the holoenzyme
aids in binding and releasing the lagging-strand template by participating in the
removal and reassembly of the sliding clamp formed by the (3 subunits.

The replisome model explains how synthesis of the leading and lagging strands is coor-
dinated. The structure of the replisome also ensures that all the components necessary for

20.6 DNA Replication Technologies 615

replication are available at the right time, in the right amount, and in the right place. Com-
plexes of proteins that function together to carry out a biochemical task are frequently called
protein machines. The replisome is an example of a protein machine, as are the bacterial fla-
gellum (Chapter 4), the ATP synthase complex (Chapter 14), the photosynthetic reaction
center (Chapter 15), and several others that are discussed in the following chapters.

20.5 Initiation and Termination
of DNA Replication

As noted earlier, DNA replication begins at a specific DNA sequence called the origin. In E.
coli , this site is called oriC , and it is located at about 10 o’clock on the genetic map of the
chromosome (Figure 20.16). The initial assembly of replisomes at oriC depends on pro-
teins that bind to this site causing local unwinding of the DNA. One of these proteins,
DnaA, is encoded by the dnaA gene that is located very close to the origin. DnaA helps reg-
ulate DNA replication by controlling the frequency of initiation. The initial RNA primers
required for leading- strand synthesis are probably made by the primosomes at the origin.

Termination of replication in E. coli occurs at the termination site ( ter ), a region
opposite the origin on the circular chromosome. This region contains DNA sequences
that are binding sites for a protein known as terminator utilization substance (Tus). The
structure of Tus bound to a single termination site is shown in Figure 20.17. Regions of
/ 3 strand lie in the major groove of DNA where the amino acid side chains make contact
with the base pairs and recognize the ter sequence. Tus prevents replication forks from
passing through the region by inhibiting the helicase activity of the replisome. The ter-
mination site also contains DNA sequences that play a role in the separation of daughter
chromosomes when DNA replication is completed.

20.6 DNA Replication Technologies

Our understanding of the basic principles of DNA replication has led to the develop-
ment of some amazing technologies that Watson and Crick could never have anticipated
in 1953. We have already encountered site-directed matagenesis (Box 6.1). In this
section we explore amplification and sequencing technologies that have transformed
biochemistry and, indeed, all biology. These technologies have produced genome
sequences of extinct species (e.g., Homo neanderthalensis) and to the discovery of the
genetic basis of many human traits and diseases.

A. The Polymerase Chain Reaction Uses DNA Polymerase
to Amplify Selected DNA Sequences

The polymerase chain reaction (PCR) is a valuable tool for amplifying a small amount of
DNA or increasing the proportion of a particular DNA sequence in a population of
mixed DNA molecules. The use of PCR technology avoids the need to take large sam-
ples of tissue in order to obtain enough DNA to manipulate for sequencing or cloning.
The polymerase chain reaction also enables the production of a large number of copies
of a gene that has not been isolated but whose sequence is known. It thus can serve as an
alternative to cloning for gene amplification.

The PCR technique is illustrated in the figure on page 621. Sequence information
from both sides of the desired locus is used to construct oligonucleotide primers that
flank the DNA sequence to be amplified. The oligonucleotide primers are complementary
to opposite strands and their 3 ' ends are oriented toward each other. The DNA from the
source (usually representing the entire DNA in a cell) is denatured by heating in the pres-
ence of excess oligonucleotides. On cooling, the primers preferentially anneal to their
complementary sites, which border the DNA sequence of interest. The primers are then
extended using a heat- stable DNA polymerase, such as Taq polymerase from the ther-
mophilic bacterium Thermus aquaticus. After one cycle of synthesis, the reaction mixture

Figure 20.17 ►

Structure of E. colilus bound to DNA. Tus binds to specific sequences at the termination site of DNA
replication. The bound protein blocks movement of the replisome. [PDB 1ECR].

▲ Protein machines. Sometimes the machine
metaphor can be taken too literally
as in this humorous cover from the Journal
Structure.

▲ Figure 20.16

Location of the origin (or/C) and terminus [ter)
of DNA replication in E. coli. dnaA is the gene
for the protein DnaA, which is required to
initiate replication. The distance between
oriC and dnaA is about 40 kb. The red ar-
rows indicate the direction of movement of
the replication forks.

616

CHAPTER 20 DNA Replication, Repair, and Recombination

is again heated to dissociate the DNA strands and cooled to reanneal the DNA with the
oligonucleotides. The primers are then extended again. In this second cycle, two of the
newly synthesized, single-stranded chains are precisely the length of the DNA between the
5' ends of the primers. The cycle is repeated many times, with reaction time and tempera-
ture carefully controlled. With each cycle, the number of DNA strands whose 5' and 3'
ends are defined by the ends of the primers increases exponentially, whereas the number
of DNA strands including sequences outside the region bordered by the primers increases
arithmetically. As a result, the desired DNA is preferentially replicated until, after 20 to 30
cycles, it makes up most of the DNA in the test tube. The target DNA sequence can then
be cloned, sequenced, or used as a probe for screening a recombinant DNA library.

B. Sequencing DNA Using Dideoxynucleotides

In 1976 Frederick Sanger developed a method for sequencing DNA enzymatically using
the Klenow fragment of E. coli DNA polymerase I. Sanger was awarded his second
Nobel Prize for this achievement (he received his first Nobel Prize for developing a
method for sequencing proteins). The advantage of using the Klenow fragment for this
type of reaction is that the enzyme lacks 5' — » 3' exonuclease activity, which could de-
grade newly synthesized DNA. However, one of the disadvantages is that the Klenow
fragment is not very processive and is easily inhibited by the presence of secondary
structure in the single-stranded DNA template. This limitation can be overcome by
adding SSB or analogous proteins, or more commonly, by using DNA polymerases from
bacteria that grow at high temperatures. Such polymerases are active at 60° to 70°C, a
temperature at which secondary structure in single-stranded DNA is unstable.

The Sanger sequencing method uses 2', 3'-dideoxynucleoside triphosphates
(ddNTPs), which differ from the deoxyribonucleotide substrates of DNA synthesis by
lacking a 3 '-hydroxyl group (see below). The dideoxyribonucleotides, which can serve
as substrates for DNA polymerase, are added to the 3' end of the growing chain. Be-
cause these nucleotides lack a 3 '-hydroxyl group, subsequent nucleotide additions can-
not take place and incorporation of a dideoxynucleotide terminates the growth of the
DNA chain. When a small amount of a particular dideoxyribonucleotide is included in
a DNA synthesis reaction, it is occasionally incorporated in place of the corresponding
dNTP, immediately terminating replication. The length of the resulting fragment of
DNA identifies the position of the nucleotide that should have been incorporated.

DNA sequencing using ddNTP molecules involves several steps (as shown on
page 622). The DNA is prepared as single-stranded molecules and mixed with a short
oligonucleotide complementary to the 3' end of the DNA to be sequenced. This oligonu-
cleotide acts as a primer for DNA synthesis catalyzed by DNA polymerase. The
oligonucleotide-primed material is split into four separate reaction tubes. Each tube re-
ceives a small amount of an a- [ 32 P] -labeled dNTP, whose radioactivity allows the newly
synthesized DNA to be visualized by autoradiography. Next, each tube receives an excess
of the four nonradioactive dNTP molecules and a small amount of one of the four
ddNTPs. For example, the A reaction tube receives an excess of nonradioactive dTTP,
dGTP, dCTP, and dATP mixed with a small amount of ddATP. DNA polymerase is then
added to the reaction mixture. As the polymerase replicates the DNA, it occasionally in-
corporates a ddATP residue instead of a dATP residue, and synthesis of the growing
DNA chain is terminated. Random incorporation of ddATP results in the production of
newly synthesized DNA fragments of different lengths, each ending with A (i.e., ddA).
The length of each fragment corresponds to the distance from the 5 '-end of the primer
to one of the adenine residues in the sequence. Adding a different dideoxyribonucleotide

°0— P — O— P — O— P — o —

► Chemical structure of a 2',3'-dideoxynucleoside
triphosphate. B represents any base.

20.6 DNA Replication Technologies 617

5 '

3 '

5 '

Heat melts DNA duplex.
Primers are added.

(1), (2)

◄ Three cycles of the polymerase chain
reaction. The sequence to be amplified
is shown in blue. (1) The duplex DNA is
melted by heating and cooled in the pres-
ence of a large excess of two primers (red
and yellow) that flank the region of interest.
(2) A heat-stable DNA polymerase catalyzes
extension of these primers, copying each
DNA strand. Successive cycles of heating
and cooling in the presence of the primers
allow the desired sequence to be repeatedly
copied until, after 20 to 30 cycles, it repre-
sents most of the DNA in the reaction
mixture.

618 CHAPTER 20 DNA Replication, Repair, and Recombination

Sanger method for sequencing DNA. ►

Addition of a small amount of a particular
dideoxynucleoside triphosphate (ddNTP) to
each reaction mixture causes DNA synthesis
to terminate when that dideoxynucleotide
is incorporated in place of the normal
nucleotide. The positions of incorporated
dideoxynucleotides, determined by the
lengths of the DNA fragments, indicate the
positions of the corresponding nucleotide
in the sequence. The fragments generated
during synthesis with each ddNTP are
separated by size using an electrophoretic
sequencing gel, and the sequence of the
DNA can be read from an autoradiograph
of the gel (as shown by the column of letters
to the right of the gel).

Single-stranded DNA
(template sequence _
unknown)

MB- T

mk g

SoBfe- c

DNA template and primer
.3'

^ 3' ^Primer

Divide into four
separate reaction tubes

dTTP, dGTP,
and dCTP

a- 32 P-dNTP
+

dTTP, dATP, dTTP, dGTP,

and dCTP and dATP

a - 32 P-dNTP
+

u- 32 P-dNTP

ddATP, dATP ddGTP, dGTP ddCTP, dCTP

dATP, dGTP,
and dCTP

a - 32 P-dNTP
+

ddTTP, dTTP

Add DNA
polymerase

Fragments end Fragments end Fragments end

ddA 3 '

in ddG

in ddC

in ddT

Separate newly synthesized
fragments from templates

Smallest®

5 '

to each reaction tube produces a different set of fragments: ddTTP produces fragments
that terminate with T, ddGTP with G, and ddCTP with C. The newly synthesized chains
from each sequencing reaction are separated from the template DNA. Finally, the mix-
tures from each sequencing reaction are subjected to electrophoresis in adjacent lanes on
a sequencing gel, where the fragments are resolved by size. The sequence of the DNA
molecule can then be read from an autoradiograph of the gel.

This technique has also been modified to allow automation for high throughput
applications like genomic sequencing. Instead of using radioactivity, automated se-
quencing relies on fluorescently labeled deoxynucleotides (four colors, one for each
base) to detect the different chain lengths. In this system the gel is “read” by a fluorime-
ter and the data are stored in a computer file. Additionally, the sequencing machine can
also provide a graphic chromatogram that shows the location and size of each fluores-
cent peak on the gel as they passed the detector.

20.7 DNA Replication in Eukaryotes 619

C. Massively Parallel DNA Sequencing by Synthesis

The automated DNA sequencing methods used to sequence the human genome have now
been largely supplanted by a variety of so-called “next generation” sequencing technolo-
gies. While using slightly different experimental approaches, these devices can all rapidly
generate millions (or even billions) of base pairs of sequence at a fraction of the cost of the
automated Sanger technology described in the previous section. As an example of this
novel approach, we describe the Illumina next- generation sequencing protocol.

In the first step, DNA (typically the entire genome) is randomly fragmented by
shearing to yield short double-stranded fragments. The ends of the fragments are enzy-
matically repaired and a single -stranded oligonucleotide primer is ligated onto each end.
Fragments of the desired length are purified from an agarose gel and then amplified
using PCR. Oligonucleotides complementary to the PCR primers are covalently attached
to the surface of a glass slide. The amplified genomic fragments are denatured into single
strands, diluted, and hybridized to the oligonucleotides on the slide.

This creates a slide where millions of individual DNA fragments are bound to the
surface. Each one is surrounded by a zone of free oligonucleotides bound to the surface.
The individual DNA fragments on the slide’s surface are then amplified in situ using a
bridging technique to yield amplification clusters that are the substrate for the
sequencing reaction.

All of the clusters of amplified DNA fragments are sequenced at the same time, in
parallel, using a mixture of the four dNTPs that have been labeled with a removable flu-
orphore (a different dye for each base) and a reversible terminator at the 3 'position (see
below). To increase the efficiency of this step, a genetically engineered mutant DNA
polymerase from the deep hydrothermal vent archeon 9°N-7 that efficiently incorpo-
rates these bulky substrates is used. The DNA sequencing primer annealed to the tem-
plate strands provides the 3' hydroxyl group and the polymerase incorporates the next
labeled nucleotide. The terminator at the 3 'position of the incoming base prevents
DNA synthesis beyond one single base. The slide is scanned using a laser-scanning con-
focal microscope to record the base that was incorporated into each growing cluster.
The reducing agent TCEP is then added removing both the dye and the terminator to
regenerate the 3' -OH. The whole cycle is then repeated. The growing DNA chains can
only increase in length via a stepwise process: one base at a time.

The relatively short sequences (less than 100 nucleotides) are not suitable for assem-
bling the genome sequence from a species that has never been sequenced before. However
for resequencing a previously sequenced genome, fast computer algorithms can align
these short “reads” with high accuracy and detect rare mutations or polymorphisms pres-
ent in the sample.

▲ Imaging clusters during the sequencing
process. Part of the image of a flow-cell with
a low density of clusters is shown. Since
each of the four deoxynucleotide bases is la-
beled with a different fluophore (each of
which fluoresces at a different wavelength),
the four separate images have been super-
imposed (after artificial coloring). After each
cycle of DNA synthesis these images provide
the raw data that reveal the last base that
was incorporated into the growing polynu-
cleotide chain.

Source: Bentley etal. (2008). Nature 456:53-59.

◄ Structure of the reversible terminator
3'-0-azidomethyl 2'-deoxythymine triphos-
phate labeled with a removable fluorophore.
Source: Bentley et al. (2008). Nature 456: 53-59.

20.7 DNA Replication in Eukaryotes

The mechanisms of DNA replication in prokaryotes and eukaryotes are fundamentally
similar. In eukaryotes as in E. coli , synthesis of the leading strand is continuous and syn-
thesis of the lagging strand is discontinuous. Furthermore, in both prokaryotes and eu-
karyotes, synthesis of the lagging strand is a stepwise process involving: primer synthe-
sis, Okazaki fragment synthesis, primer hydrolysis, and gap filling by a polymerase.
Eukaryotic primase, like prokaryotic primase, synthesizes a short primer once every
second on the lagging-strand template. However, because the replication fork moves
more slowly in eukaryotes, each Okazaki fragment is only about 100 to 200 nucleotide

620

CHAPTER 20 DNA Replication, Repair, and Recombination

Table 20.2 Eukaryotic DNA polymerases

DNA polymerase

Activities

Role

Polymerase

Primase

3' — >5' Exonuclease 0

Primer synthesis
Repair

Polymerase

Repair

Polymerase

3' — >5' Exonuclease

Mitochondrial DNA replication

Polymerase

3' — >5' Exonuclease

Leading- and lagging-strand synthesis
Repair

Polymerase

3' — >5' Exonuclease

5' — > 3' Exonuclease

Repair

Gap filling on lagging strand

°Polymerase a3’ — > 5' exonuclease activity is not detectable in all species.

residues long, considerably shorter than in prokaryotes. Interestingly, eukaryotic DNA
primase does not share significant sequence similarity with the E. coli enzyme nor does
eukaryotic primase contain some of the classical structural landmarks of DNA poly-
merases such as the “fingers” or “thumb” domains (Figure 20.12). This lack of homol-
ogy suggests that the capacity to synthesize an RNA primer for DNA initiation may have
evolved independently at least twice.

Most eukaryotic cells contain at least five different DNA polymerases: a , /3, y, d,
and s (Table 20.2). DNA polymerases a , d, and £ are responsible for the chain elonga-
tion reactions of DNA replication and for some repair reactions. DNA polymerase /3 is a
DNA repair enzyme found in the nucleus and DNA polymerase y plays a role in repli-
cating mitochondrial DNA. A sixth DNA polymerase is responsible for replicating DNA
in chloroplasts.

DNA polymerase d catalyzes synthesis of the leading strand at the replication fork.
This enzyme is composed of two subunits the larger of which contains the polymerase
active site. The enzyme also has 3' — > 5' exonuclease activity. DNA replication in eu-
karyotic cells is extremely accurate. The low error rate indicates that DNA replication in
eukaryotes includes an efficient proofreading step.

DNA polymerase a and DNA polymerase d cooperate in lagging strand synthesis.
DNA polymerase a is a multimeric protein that contains both DNA polymerase and
RNA primase activity. The primer made by DNA polymerase a consists of a short
stretch of RNA followed by DNA. This two part primer is extended by DNA polymerase
d to complete an Okazaki fragment.

DNA polymerase £ is a large, multimeric protein. The largest polypeptide chain in-
cludes polymerase activity and 3' —> 5' proofreading exonuclease activity. Like its func-
tional counterpart in E. coli (DNA polymerase I), DNA polymerase £ probably acts as a
repair enzyme and also fills gaps between Okazaki fragments.

Several accessory proteins are associated with the replication fork in eukaryotes.
These proteins function like some of the proteins in the bacterial replisome. For exam-
ple, PCNA (proliferating cell nuclear antigen) forms a structure that resembles the
/3-subunit sliding clamp of E. coli DNA polymerase III (Figure 20.7). The accessory pro-
tein RPC (replication factor C) is structurally, functionally, and evolutionarily related to
the y complex of DNA polymerase III. Another protein, called RPA (replication factor A),
is the eukaryotic equivalent of prokaryotic SSB. In addition, the eukaryotic replication
machine includes helicases that unwind DNA at the replication fork.

Each eukaryotic chromosome contains many origins of replication (Section 20.1).
The largest chromosome of the fruit fly Drosophila melanogaster , for example, contains
about 6000 replication forks implying that there are at least 3000 origins. As replication
proceeds bidirectionally from each origin the forks move toward one another, merging
to form bubbles of ever increasing size (Figure 20.4). Due to the large number of ori-
gins, the larger chromosomes of eukaryotes can still be replicated in less than one hour
even though the rate of individual fork movement is much slower than in prokaryotes.

20.7 DNA Replication in Eukaryotes 621

The eukaryotic cell division cycle coordinates
DNA replication and mitosis. DNA replication
occurs exclusively during the synthesis, or
S-phase of the cell cycle. There are two gap ;
or G, phases where a cell grows prior to di-
viding in the mitosis, or M-phase.

◄ Figure 20.18

DNA replication in all cells occurs within the context of the cells programmed cell di-
vision cycle. This cell cycle is a highly regulated progression through a series of dependent
steps that at a minimum accomplishes two goals: (1) it faithfully duplicates all of the DNA
in a cell to produce exactly two copies of each chromosome, and (2) it precisely segregates
one copy of each replicated chromosome into one of the two daughter cells. In eukaryotic
cells chromosomal segregation occurs at mitosis and this stage is called the mitotic phase,
or M-phase (Figure 20.18). The step where DNA is synthesized is called S-phase. The inter-
phase (resting) stage between mitosis and the next round of DNA replication is called Gl.
There may be a G2 stage between the end of DNA replication and the beginning of mitosis.

Eukaryotic DNA replication origins must be used once, and only once, during S-
phase of each cell cycle. We are beginning to understand some of the key players that or-
chestrate this process. At the end of the previous M-phase and during the subsequent Gl-
phase, each functional ori becomes an assembly site for a conserved multiprotein complex
named ORC (origin recognition complex). As the cell progresses through Gl each ORC
stimulates the formation of a prereplication complex (pre-RC) that includes a helicase.
The pre-RC remains poised until the activity of an S-phase protein kinase (SPK) drops to
a critical threshold, whereby the initiation complex recruits waiting replisomes and the
origin is said to “fire.” The two replication forks are then launched along the chromosome
in opposite directions. When SPK activity is high it prevents any new pre-RCs from load-
ing onto the origins, thus preventing multiple rounds of initiation. SPK is proteolytically
cleaved at the beginning of the mitotic phase allowing ORC proteins to bind to the origins
waiting on each daughter chromosome beginning late in M-phase.

Eukaryotic replication origins do not all fire simultaneously at the beginning of S-
phase. Instead, transcribed, or active, regions of a cell’s genome tend to be replicated
earlier during S-phase while the origins located in quiescent, or repressed, regions of the
genome tend to be replicated later in S-phase. It remains to be determined whether this
differential timing of replication actually depends on transcription or just reflects that
“open” chromatin permits ORC to locate replication origins.

The differences between eukaryotic and prokaryotic DNA replication arise not only
from the larger size of the eukaryotic genome but also from the packaging of eukaryotic
DNA into chromatin. The binding of DNA to histones and its packaging into nucleo-
somes, (Section 19.5), is thought to be responsible in part for the slower movement of

622 CHAPTER 20 DNA Replication, Repair, and Recombination

Figure 20.19 ►

Photodimerization of adjacent deoxythymidy-
late residues. Ultraviolet light causes the
bases to dimerize, thus distorting the struc-
ture of DNA. For clarity, only a single strand
of DNA is shown.

0 =

h 3 c

H O

Figure 20.20 ►

Repair of thymine dimers by DNA photolyase.

In the presence of visible
light, the enzyme catalyzes
chemical cleavage of the
dimer, thereby restoring
normal base pairing and
repairing the DNA.

20.8 Repair of Damaged DNA

623

the replication fork in eukaryotes. Eukaryotic DNA replication occurs with concomi-
tant synthesis of histones; the number of histones doubles with each round of DNA
replication. Histone duplication and DNA replication involve different enzymes acting
in different parts of the cell yet both occur at about the same rate. It appears that exist-
ing histones remain bound to DNA during replication and that newly synthesized his-
tones bind to DNA behind the replication fork shortly after synthesis of the new
strands.

20.8 Repair of Damaged DNA

DNA is the only cellular macromolecule that can be repaired. This is probably because
the cost to the organism of mutated or damaged DNA far outweighs the energy spent to
repair the defect. Repairing other macromolecules is not profitable; for example, little is
lost when a defective protein forms as a result of a translation error because the protein
is simply replaced by a new, functional protein. When DNA is damaged, however, the
entire organism may be in jeopardy if the instructions for synthesizing a critical mole-
cule are altered. In single-celled organisms, damage to a gene encoding an essential pro-
tein may kill the organism. Even in multicellular organisms, the accumulation of defects
in DNA over time can lead to progressive loss of cellular functions or to deregulated
growth such as that seen in cancer cells.

There are several types of DNA damage such as base modifications, nucleotide
deletions or insertions, cross-linking of DNA strands, and breakage of the phosphodi-
ester backbone. While some DNA damage is the result of environmental agents (e.g.,
chemicals or radiation) most DNA damage is the result of errors made during DNA
replication. Severe damage may be lethal but much of the damage that occurs in vivo is
repaired. Many modified nucleotides, as well as mismatched bases that escape the
proofreading mechanism of DNA polymerase, are recognized by specific repair en-
zymes that continually scan DNA in order to detect alterations. Some of the lesions are
fixed by direct repair, a process that does not require breaking the phosphodiester back-
bone of DNA. Other repairs require more extensive work.

DNA repair mechanisms protect individual cells as well as subsequent generations.
In single- celled organisms, whether prokaryotes or eukaryotes, DNA damage that is not
repaired may become a mutation that is passed directly to the daughter cells following
DNA replication and cell division. In multicellular organisms, mutations can be passed
on to the next generation only if they occur in the germ line. Germ line mutations may
have no noticeable effect on the organism that contains them but may have profound
effects on the progeny, especially if the mutated genes are important in development.
When mutations occur in somatic cells however, while the defects are not transmissible,
they can sometimes lead to unrestricted cell growth, or cancer. In spite of the accuracy
of DNA replication and the efficiency of repair, the average human accumulates about
130 new mutations every generation. Most of these mutations are neutral and this leads
to a huge amount of variation in human populations. It is this variation that makes pos-
sible the identification of individuals by DNA fingerprinting.

A. Repair after Photodimerization: An Example of Direct Repair

Double-helical DNA is susceptible to damage by ultraviolet (UV) light. The most com-
mon UV light-induced damage is dimerization of adjacent pyrimidines in a DNA
strand. This process is an example of photodimerization. The most common dimers
form between adjacent thymines (Figure 20.19). DNA replication cannot occur in the
presence of pyrimidine dimers because they distort the template strand. Therefore,
removal of pyrimidine dimers is essential for survival.

Many organisms can repair thymine dimer damage using direct repair (notably,
humans and all placental mammals lack this repair mechanism — see below). The sim-
plest repair process begins when an enzyme known as DNA photolyase binds to the dis-
torted double helix at the site of the thymine dimer (Figure 20.20). As the DNA-enzyme
complex absorbs visible light, the dimer is cleared. The photolyase then dissociates
from the repaired DNA and normal A/T base pairs re-form. This process is called photo
reactivation; it’s an example of direct repair.

624 CHAPTER 20 DNA Replication, Repair, and Recombination

Site of damage

Excision-repair enzymes
detect damaged DNA. An
endonuclease nicks the DNA
backbone on both sides of the
damage.

A helicase or exonuclease
removes the damaged DNA,
leaving a gap.

$OCsS%o$o<

DNA polymerase fills the
gap.

The remaining nick is
sealed by DNA ligase.

▲ Figure 20.21

General excision-repair pathway.

B. Excision Repair

Other forms of ionizing radiation and naturally occurring chemicals
can damage DNA. Some compounds, including acids and oxidizing
agents, can modify DNA by alkylation, methylation, or deamination.
DNA is also susceptible to spontaneous loss of heterocyclic bases, a
process known as depurination or depyrimidization. Many of these
defects can be repaired by a general excision repair pathway whose
overall features are similar in all organisms. The pathway begins
when an endonuclease recognizes distorted, damaged DNA and
cleaves on both sides of the lesion releasing an oligonucleotide con-
taining 12 to 13 residues. This cleavage is catalyzed by the UvrABC
enzyme in E. coli. Removal of the DNA oligonucleotide may require
helicase activity that is often a component of the excision repair en-
zyme complex. The result is a single-stranded gap. The gap is then
filled in by the action of DNA polymerase I in prokaryotes or repair
DNA polymerases in eukaryotes. The nick is sealed by DNA ligase
(Figure 20.21).

The UvrABC endonuclease also recognizes pyrimidine dimers
and modified bases that distort the double helix (this is how thymine
dimers are repaired in humans). Other excision-repair enzymes rec-
ognize DNA damaged by hydrolytic deamination of adenine, cyto-
sine, or guanine. (Thymine is not subject to deamination because it
does not have an amino group.) The deaminated bases can form in-
correct base pairs resulting in the incorporation of incorrect bases
during the next round of replication. Spontaneous deamination of
cytosine is one of the most common types of DNA damage because
the product of deamination is uracil that easily forms a base pair with
adenine in the next round of replication (Figure 20.22).

Enzymes called DNA glycosylases remove deaminated bases and
some other modified bases by catalyzing hydrolysis of the N-glycosidic
bonds that link the modified bases to the sugars. Let’s look at the re-
pair of deaminated cytosine. Repair begins when the enzyme uracil
N-glycosylase removes the uracil produced by deamination. The en-
zyme recognizes and binds to the incorrect U/G base pair and flips the
uracil base outward, positioning the /3-N-glycosidic bond in the active
site of the enzyme where it is cleaved from the sugar residue (Figure
20.23). Next, an endonuclease recognizes the site where the base is
missing and removes the deoxyribose phosphate, leaving a single-nucleotide gap in the
duplex DNA. The endonuclease is called an AP-endonuclease because it recognizes
apurinic and apyrimidinic sites (AP sites). Some specific DNA glycosylases are bifunc-
tional enzymes with both glycosylase and AP-endonuclease activities in the same
polypeptide chain. Excision repair enzymes with exonuclease activity often extend the
gap produced by the endonuclease. In prokaryotes, DNA polymerase I binds to the ex-
posed 3' end of DNA and fills in the gap. Finally, the strand is sealed by DNA ligase. The
steps of the excision repair pathway are summarized in Figure 20.24.

Whereas deamination of adenine or guanine is rare, deamination of cytosine is
fairly common and would give rise to large numbers of mutations were it not for the re-
placement of uracil with thymine in DNA. (Recall that thymine is simply 5-methylu-
racil.) If uracil were normally found in DNA, as it is in RNA, it would be impossible to
distinguish between a correct uridylate residue and one arising from the deamination of
cytosine. However, since uracil is not one of the bases in DNA, damage arising from cy-
tosine deamination can be recognized and repaired. Thus, the presence of thymine in
DNA increases the stability of genetic information.

•Ml

20.8 Repair of Damaged DNA 625

Uracil produced by
deamination of cytosine

^ Uracil is recognized by

C uracil A/-glycosylase, which

hydrolyzes the A/-glycosidic
APsite bond, yielding an AP site.

I An endonuclease recognizes
the AP site, cleaves the sugar-
phosphate backbone, and
removes the deoxyribose
phosphate.

I The resulting single-
nucleotide gap is filled
by DNA polymerase I,
and the nick is sealed
by DNA ligase.

Amino group

HC^

Cytosine

H 2 0

nh 3

Hydrolytic

deamination

HC"

"NH

Uracil

▲ Figure 20.22

Hydrolytic deamination of cytosine. Deamination
of cytosine produces uracil, which pairs with
adenine rather than guanine.

▲ Figure 20.23

Uracil /V-glycosylase from human mitochon-
dria. The enzyme is bound to a uracil-
containing nucleotide (green) that has been
flipped out of the stacked region of double-
stranded DNA. [PDB 1EMH].

▲ Figure 20.24

Repair of damage resulting from the deamination of cytosine.

626

CHAPTER 20 DNA Replication, Repair, and Recombination

BOX 20.1 THE PROBLEM WITH METHYLCYTOSINE

5-Methylcytosine is common in eukaryotic DNA (Section 18.7). Deamination of
5-methylcytosine produces thymidine giving rise to a T opposite a G in damaged
DNA. Repair enzymes cannot recognize which of these bases is incorrect, so the
“repair” often results in a T:A base pair. This will also happen if the damaged DNA
is replicated before it can be repaired. The cytosines at CG sites are preferentially
methylated in mammalian genomes. Frequent loss of the cytosines by deamination
of 5-methylcytosine has led to underrepresentation of CG sequences relative to TG,
AG, and GG.

20.9 Homologous Recombination

Recombination is any event that results in the exchange or transfer of pieces of DNA
from one chromosome to another or within a chromosome. Most recombinations are
examples of homologous recombination because they occur between pieces of DNA that
have closely related sequences. Exchanges between paired chromosomes during meiosis
are examples of homologous recombination. Recombination between unrelated se-
quences is called nonhomologous recombination. Transposons are mobile genetic elements
that jump from chromosome to chromosome by taking advantage of nonhomologous
recombination mechanisms. Recombination between DNA molecules also occurs when
bacteriophages integrate into host chromosomes. When recombination occurs at a spe-
cific location it is called site specific recombination.

Mutation creates new genetic variation in a population and recombination is a
mechanism that creates different combinations of mutations in a genome. Most species
have some mechanism for exchanging information between individual organisms.
Prokaryotes usually contain only a single copy of their genome (i.e., they are haploid),
so this exchange requires recombination. Some eukaryotes are also haploid but most are
diploid, having two sets of chromosomes, one contributed by each parent. Genetic re-
combination in diploids mixes the genes on the chromosomes contributed by each par-
ent so that subsequent generations receive very different combinations of genes. None
of your childrens chromosomes, for example, will be the same as yours and none of
yours are the same as those of your parents. (Although this mixing of alleles is an im-
portant consequence of recombination, it is not likely to be the reason why recombina-
tion mechanisms evolved in the first place. The problem of why sex evolved is one of the
most difficult problems in biology.)

Recombination occurs by many different mechanisms. Many of the proteins and
enzymes that participate in recombination reactions are also involved in DNA repair
reactions illustrating the close connection between repair and recombination. In this
section, we briefly describe the Holliday model of general recombination — a type of
recombination that seems to occur in many species.

A. The Holliday Model of General Recombination

Homologous recombination begins with the introduction of either single-stranded or
double- stranded breaks into DNA molecules. Recombination involving single-stranded
breaks is often called general recombination. Recombination involving double-stranded
breaks is not discussed here, although it is an important mechanism of recombination
in some species.

Consider general recombination between two linear chromosomes as an example
of recombination in prokaryotes. The exchange of information between the molecules
begins with the alignment of homologous DNA sequences. Next, single- stranded nicks
are introduced in the homologous regions and single strands exchange in a process
called strand invasion. The resulting structure contains a region of strand crossover and

20.9 Homologous Recombination 627

Homologous chromosomes
pair and are nicked.

Strand invasion occurs.

Lower strand
rotates 180°.

Left ends
rotate 180°

DNA is cleaved at
crossover point and

◄ Figure 20.25

Holliday model of general recombination.

Nicks are introduced into a homologous
region of each molecule. Subsequent strand
invasion, DNA cleavage at the crossover
junction, and sealing of nicked strands
result in exchange of the ends of the
chromosomes.

▲ Asexual Daphnia

is known as a Holliday junction after Robin Holliday who first proposed it in 1964
(Figure 20.25).

The chromosomes can be separated at this stage by cleaving the two invading
strands at the crossover point. It is important to realize that the ends of the homologous
DNA molecules can rotate generating different conformations of the Holliday junction.
Rotation followed by cleavage produces two chromosomes that have exchanged ends as
shown in Figure 20.25. Recombination in many different organisms probably occurs by
a mechanism similar to the one shown in Figure 20.25.

B. Recombination in f. coli

One of the first steps in recombination is the generation of single-stranded DNA with a
free 3' end. In E. coli , this step is carried out by RecBCD endonuclease, an enzyme with
subunits that are encoded by three genes (recB, recC, and recD) whose products have
long been known to play a role in recombination. RecBCD binds to DNA and cleaves

▲ Male Drosophila melanogaster (no meiotic
recombination)

628 CHAPTER 20 DNA Replication, Repair, and Recombination

Meiotic chisasmata ►

Source: © 2008 Sinauer Associates Sadava, D. et al.
Life: The Science of Biology, 8th ed. (Sunderland,
MA: Sinauer Associates and W. H. Freeman &
Company), 198

RecA

RecA-coated strand
binds to homologous
w double-stranded DNA.

Homologous DNA

Triple-stranded

intermediate

Strand invasion and
displacement occur.

Branch migration
extends the region
of exchange.

Exchange is
completed.

▲ Figure 20.26

Strand exchange catalyzed by RecA.

Homologous

chromosomes

Chiasmata

Centromeres

one of the strands. It then unwinds the DNA in a process coupled to ATP hydroly-
sis generating single-stranded DNA with a 3' terminus.

Strand exchange during recombination begins when the single-stranded
DNA invades the double helix of a neighboring DNA molecule. Strand exchange
is not a thermodynamically favorable event — the invasion must be assisted by pro-
teins that promote recombination and repair. RecA is the prototypical strand ex-
change protein. It is essential for homologous recombination and for some forms
of repair. The protein functions as a monomer that binds cooperatively to single-
stranded DNA such as the single- stranded tails produced by the action of RecBCD.
Each RecA monomer covers about five nucleotide residues and each successive
monomer binds to the opposite side of the DNA strand.

One of the key roles of RecA in recombination is to recognize regions of se-
quence similarity. RecA promotes the formation of a triple-stranded intermediate
between the RecA-coated single strand and a highly similar region of double-
stranded DNA. RecA then catalyzes strand exchange in which the single strand dis-
places the corresponding strand from the double helix.

Strand exchange takes place in two steps: strand invasion, followed by branch
migration (Figure 20.26). Both the single- stranded and the double-stranded DNA
are in an extended conformation during the exchange reaction. The strands must
rotate around each other, a process that is presumably aided by topoisomerases.
Strand exchange is a slow process despite the fact that no covalent bonds are bro-
ken. (A “slow” process in biochemistry is one that takes several minutes.)

RecA can also promote strand invasion between two aligned, double-stranded
DNA molecules. Both molecules must contain single-stranded tails bound to RecA.
The tails wind around the corresponding complementary strands in the homo-
logue. This exchange gives rise to a Holliday junction such as the one shown in
Figure 20.25. Subsequent branch migration can extend the region of strand ex-
change. Branch migration can continue even after RecA dissociates from the re-
combination intermediate.

Branch migration at the double-stranded version of a Holliday junction is
driven by a remarkable protein machine found in all species. The bacterial version
is made up of RuvA and RuvB subunits. These proteins bind to the junction and

◄ Bacterial conjugation (or sex).

20.9 Homologous Recombination 629

promote branch migration as shown in the schematic diagram (Figure 20.27). The two
DNA helices are separated when RuvC binds to the Holliday junction and cleaves the
crossover strands.

RuvA and RuvB form a complex consisting of four RuvA subunits bound to the
Holliday junction and two hexameric rings of RuvB subunits that surround two of the
DNA strands (Figure 20.28). The RuvB component is similar to the sliding clamps
discussed in the section on DNA replication (Section 20. 2B) and it drives branch mi-
gration by pulling the strands through the RuvA/Holliday junction complex in a reac-
tion coupled to ATP hydrolysis (Figure 20.29). The rate of RuvAB- mediated branch mi-
gration is about 100, 000 bp per second — significantly faster than strand invasion.

RuvC catalyzes cleavage of the crossover strands to resolve Holliday junctions. Two
types of recombinant molecules are produced as a result of this cleavage: those in which
only single strands are exchanged and those in which the ends of the chromosome have
been swapped (Figure 20.25).

C. Recombination Can Be a Form of Repair

▲ RecBCD bound to DNA showing separation of
strands. [PDB 3K70]

Since natural selection works predominantly at the level of individual organisms it is
difficult to see why recombination would have evolved unless it affected survival of the
individual. Recombination enzymes probably evolved because they play a role in DNA
repair, which confers a selective advantage. For example, severe lesions in DNA are
bypassed during DNA replication, leaving a daughter strand with a single-stranded
region. RecA-mediated strand exchange between the homologous daughter chromo-
somes allows the intact strand from one daughter molecule to act as a template for
repairing the broken strand of the other daughter molecule.

Recombination also creates new combinations of genes on a chromosome and this
may be an added bonus for the population and its chances for evolutionary survival.
More than 100 E. coli genes are required for recombination and repair, and there are
twice as many in most eukaryotes.

Most, if not all, of the genes used in recombination play some role in repair as well.
Mutations in several human genes give rise to rare genetic defects that result from defi-
ciencies in DNA repair and/or recombination. For example, xeroderma pigmentosum is
a hereditary disease associated with extreme sensitivity to ultraviolet light and increased
frequency of skin cancer. Excision repair is defective in patients with this disease but the
phenotype can be due to mutations in at least eight different genes. One of these genes
encodes a DNA glycosylase with AP- endonuclease activity. Other affected genes include
some that encode helicases that are required for both repair and recombination.

Many other genetic defects related to deficiencies in repair and recombination have
not been well characterized. Some of them are responsible for increased incidences of
cancer in affected patients.

ATP

ADP+P;

RuvAB promotes
branch migration.

RuvC

RuvC binds to the
Holliday junction
and cleaves the
crossover strands.

RuvB RuvA RuvB

▲ Figure 20.27

Action of Ruv proteins at Holliday junctions.

RuvAB promotes branch migration in a reac-
tion coupled to ATP hydrolysis. RuvC cleaves
Holliday junctions. Two types of recombi-
nant molecules can be generated in this
reaction.

◄ Figure 20.28

Model of RuvA and RuvB bound to a Holliday
junction.

630 CHAPTER 20 DNA Replication, Repair, and Recombination

Junction binding

RuvA

Branch migration

Resolution

▲ Figure 20.29

Branch migration and resolution. [Adapted from Rafferty, J. B., et al. (1996). Crystal structure of DNA recombination protein RuvA and a model for its
binding to the Holliday junction. Science 274:415-421.]

BOX 20.2 MOLECULAR LINKS BETWEEN DNA REPAIR AND BREAST CANCER

About 180,000 women are diagnosed with breast cancer
every year in North America. Approximately one-fifth of
these new cases have a familial or genetic component and
one-third of these, or 12,000 cases, are due to mutations in
one of the two genes named BRCA1 or BRCA2 that encode
proteins by the same name.

Both of these proteins are required for normal recombi-
national repair of double strand breaks (DSB). BRCA2 forms
a complex with the eukaryotic RecA homologue RAD51.
BRCA2 also binds specifically to BRCA1 to form a heterotrimer.
Following exposure to ionizing radiation, these three DNA
repair proteins are found localized to discrete sites, or foci,
inside the interphase nuclei (see figure). These foci are the
sites where the proteins are repairing double strand breaks.
The BRCA proteins are so vital that cells become susceptible
to damage if either copy of the gene is damaged. When one
or both copies of the BRCA1 or BRCA2 genes are defective,
the capacity to repair DSBs is compromised leading eventu-
ally to a higher frequency of mutations. Some of these new
mutations may allow the cell to escape from the rigorous
constraints imposed by the eukaryotic cell cycle, eventually
leading to cancer. The BRCA proteins function as sentinels
by continually monitoring the genome to identify and cor-
rect potential mutagenic lesions. In fact, some humans with a
rare autosomal recessive disease called Fanconi’s Anemia
(FA) have an increased sensitivity to several mutagenic com-
pounds and a genetic predisposition to many different types

of cancers. It has been shown that FA patients are affected in
one of seven different genes that are presumably important
for DNA repair. One of these genes is BRCA2 , underscoring
its essential role in the repair process.

■ *

' * ; V. V'

* *

- -.V

■ V , V * * ' -

•M. #5*

•

V < V

- V %

#•'4 / * ", ' ’

, / * . *' J f A*

▲ Ionizing radiation induces nuclear foci of the DNA repair protein
BRCA1. Energetic y-rays can induce double-stranded breaks in
DNA and trigger DNA repair. This tissue culture cell nucleus was ex-
posed to IR and then treated with antibodies that recognize BRCA1
(stained green).

Problems 631

Summary

1. DNA replication is semiconservative; each strand of DNA serves
as the template for synthesis of a complementary strand. The
products of replication are two double- stranded daughter mole-
cules consisting of one parental strand and one newly synthesized
strand. DNA replication is bidirectional, proceeding in both di-
rections from an origin in replication.

2. DNA polymerases add nucleotides to a growing DNA chain by
catalyzing nucleotidyl- group-transfer reactions. DNA synthesis
proceeds in the 5' — » 3' direction. Errors in DNA synthesis are re-
moved by the 3' —> 5' exonuclease activity of the polymerase.
Some DNA polymerases contain an additional 5' — » 3' exo-
nuclease activity.

3. The leading strand of DNA is synthesized continuously but the
lagging strand is synthesized discontinuously producing Okazaki
fragments. Synthesis of the leading strand and of each Okazaki
fragment begins with an RNA primer. In E. coli , the primer is re-
moved and replaced with DNA by the action of DNA polymerase I.
The action of DNA ligase joins the separate fragments of the lag-
ging strand.

4. The replisome is a complex protein machine that is assembled at
the replication fork. The replisome contains two DNA polymerase
molecules plus additional proteins such as helicase and primase.

5. Assembly of the replisome ensures simultaneous synthesis of two
strands of DNA. In E. coli , a helicase unwinds the parental DNA
and SSB binds to the single strands. The lagging- strand template
is looped through the replisome so that the synthesis of both strands
proceeds in the same direction as replication fork movement.

Because it is part of the replisome, DNA polymerase is highly
processive.

6. Initiation of DNA replication occurs at specific DNA sequences
(e.g., oriC in E. coli) and depends on the presence of additional
proteins. In bacteria, termination of DNA replication also occurs
at specific sites and requires additional proteins.

7. Several new technologies such as PCR and DNA sequencing are
based on an understanding of DNA replication.

8. Eukaryotic DNA replication resembles prokaryotic DNA replica-
tion except that eukaryotic chromosomes contain multiple ori-
gins of replication and eukaryotic Okazaki fragments are smaller.
The slower movement of the replication fork in eukaryotes than
in prokaryotes is due to the presence of nucleosomes.

9. DNA damaged by radiation or chemical agents can be repaired by
direct- repair mechanisms or by a general excision- repair pathway.
Excision-repair mechanisms also remove misincorporated nu-
cleotides. Specific enzymes recognize damaged or misincorpo-
rated nucleotides.

10. Recombination can occur when a single strand of DNA exchanges
with a homologous strand in double-stranded DNA producing a
Holliday junction. Strand invasion is promoted by RecA in E. coli.
Branch migration and resolution of Holliday junctions are cat-
alyzed by RuvABC in E. coli.

11. Repair and recombination are similar processes and use many of
the same enzymes. Defects in human genes required for repair
and recombination cause sensitivity to ultraviolet light and in-
creased risks of cancer.

Problems

1. The chromosome of a certain bacterium is a circular, double-
stranded DNA molecule of 5.2 X 10 6 base pairs. The chromosome
contains one origin of replication and the rate of replication-fork
movement is 1000 nucleotides per second.

(a) Calculate the time required to replicate the chromosome.

(b) Explain how the bacterial generation time can be as short as
25 minutes under extremely favorable conditions.

2. In many DNA viruses the viral genes can be divided into two
nonoverlapping groups: early genes, whose products can be de-
tected prior to replication of the viral genome; and late genes,
whose products accumulate in the infected cell after replication of
the viral genome. Some viruses, like bacteriophage T4 and T7, en-
code their own DNA polymerase enzymes. Would you expect the
gene for T4 DNA polymerase to be in the early or late class? Why?

3. (a) Why does the addition of SSB to sequencing reactions often

increase the yield of DNA?

(b) What is the advantage of carrying out sequencing reactions at
65°C using a DNA polymerase isolated from bacteria that
grow at high temperatures?

4. How does the use of an RNA primer rather than a DNA primer
affect the fidelity of DNA replication in E. colP.

5. Both strands of DNA are synthesized in the 5' — » 3' direction.

(a) Draw a hypothetical reaction mechanism for synthesis of
DNA in the 3' — > 5' direction using a 5'-dNTP and a grow-
ing chain with a 5 '-triphosphate group.

(b) How would DNA synthesis be affected if the hypothetical en-
zyme had proofreading activity?

6. Ciprofloxacin is an antimicrobial used in the treatment of a wide
variety of bacterial infections. One of the targets of ciprofloxacin
in E. coli is topoisomerase II. Explain why the inhibition of topoi-
somerase II is an effective target to treat infections by E. coli.

7. The entire genome of the fruit fly D. melanogaster consists of
1.65 X 10 8 bp. If replication at a single replication fork occurs at
the rate of 30 bp per second, calculate the minimum time re-
quired to replicate the entire genome if replication were initiated

(a) at a single bidirectional origin

(b) at 2000 bidirectional origins

(c) In the early embryo, replication can require as few as 5 min-
utes. What is the minimum number of origins necessary to
account for this replication time?

8. Ethyl methane sulfonate (EMS) is a reactive alkylating agent that
ethylates the 0-6 residue of guanine in DNA. If this modified G is
not excised and replaced with a normal G, what would be the out-
come of one round of DNA replication?

9. Why do cells exposed to visible light following irradiation with
ultraviolet light have a greater survival rate than cells kept in the
dark after irradiation with ultraviolet light?

10 . E. coli uses several mechanisms to prevent the incorporation of
the base uracil into DNA. First, the enzyme dUTPase, encoded by the
dut gene, degrades dUTP. Second, the enzyme uracil N-glycosylase,

632 CHAPTER 20 DNA Replication, Repair, and Recombination

encoded by the ung gene, removes uracils that have found their
way into DNA. The resulting apyrimidinic sites have to be repaired.

(a) If we examine the DNA from a strain carrying a mutation in
the dut gene, what will we find?

(b) What if we examine the DNA from a strain in which both the
dut and ung genes are mutated?

11. Explain why uracil N-glycosylase cannot repair the damage when
5-methylcytosine is deaminated to thymine.

12. Why are high rates of mutation observed in regions of DNA that
contain methylcytosine?

13. Explain why the overall error rate for DNA replication in E. coli is
approximately 10~ 9 although the rate of misincorporation by the
replisome is about 10 -5 .

15. Describe two methods that can be used to repair pyrimidine
dimers in E. coli.

16. Damage to a single strand of DNA is readily repaired through a
variety of mechanisms while damage to bases on both strands of
DNA is more difficult for the cell to repair. Explain.

17. Why does homologous recombination occur only between DNAs
with identical, or almost identical, sequences?

18. Why are two different DNA polymerase enzymes required to
replicate the E. coli chromosome?

Selected Readings

General

Adams, R. L. P., Knowler, J. T., and Leader, D. P.
(1992). The Biochemistry of the Nucleic Acids, 1 1th
ed. (New York: Chapman and Hall).

Aladjem, M. I. (2007). Replication in context: dy-
namic regulation of DNA replication patterns in
metazoans. Nat. Rev. Genet. 8:588-600.

Bentley, D. R., et al. (2008). Accurate whole human
genome sequencing using reversible terminator
chemistry. Nature 456: 53-59.

Kornberg, A., and Baker, T. (1992). DNA Replica-
tion , 2nd ed. (New York: W. H. Freeman).

DNA Replication

Beese, L. S., Derbyshire, V., and Steitz, T. A.

(1993). Structure of DNA polymerase I Klenow
fragment bound to duplex DNA. Science 260:
352-355.

Bell, S. P. (2002). The origin recognition complex:
from simple origins to complex functions. Genes &
Devel. 16:659-672.

Davey, M. J., Jeruzalmi, D., Kuriyan, J., and
O’Donnell, M. (2002). Motors and switches:

AAA + machines within the replisome. Nat. Rev.
Mol. Cell Biol. 3:1-10.

Gilbert, D. M. (2001). Making sense of eukaryotic
DNA replication origins. Science 294:96-100.

Keck, J. L., and Berger, J. M. (2001). Primus inter
pares (First among equals). Nat. Struct. Biol. 8:2-4.

Kong, X.-P., Onrust, R., O’Donnell, M., and Kuriyan,
J. (1992). Three-dimensional structure of the /3
subunit of E. coli DNA polymerase III holoen-
zyme: a sliding DNA clamp. Cell 69:425-437.

Kunkel, T. A., and Bebenek, K. (2000). DNA repli-
cation fidelity. Annu. Rev. Biochem. 69:497-529.

Marians, K. J. (1992). Prokaryotic DNA replica-
tion. Annu. Rev. Biochem. 61:673-719.

McHenry, C. S. (1991). DNA polymerase III
holoenzyme. /. Biol. Chem. 266:19127-19130.

Meselson, M., and Stahl, F. W. (1958). The replica-
tion of DNA in Escherichia coli. Proc. Natl. Acad.
Sci. USA 44:671-682.

Radman, M. (1998). DNA replication: one strand
may be more equal. Proc. Natl. Acad. Sci. USA
95:9718-9719.

Waga, S., and Stillman, B. (1998). The DNA repli-
cation fork in eukaryotic cells. Annu. Rev.

Biochem. 67:721-751.

Wake, R. G., and King, G. F. (1997). A tale of two
terminators: crystal structures sharpen the debate
on DNA replication fork arrest mechanisms.
Structure 5:1-5.

Wyman, C., and Botchan, M. (1995). A familiar
ring to DNA polymerase processivity. Curr. Biol.
5:334-337.

DNA Repair

Echols, H., and Goodman, M. F. (1991). Fidelity
mechanisms in DNA replication. Annu. Rev.
Biochem. 60:477-511.

Hanawalt, P. C. and Spivak, G. (2008). Transcrip-
tion-coupled DNA repair: two decades of progress
and surprises. Nat. Rev. Mol. Cell. Biol. 9:958-970.

Kogoma, T. (1997). Stable DNA replication: inter-
play between DNA replication, homologous re-
combination, and transcription. Microbiol. Mol.
Biol. Rev. 61:212-238.

McCullough, A. K., Dodson, M. L., and Lloyd, R. S.
(1999). Initiation of base excision repair: glycosy-
lase mechanisms and structures. Annu. Rev.
Biochem. 68:255-285.

Mol, C. D., Parikh, S. S., Putnam, C. D., Lo, T. P., and
Taylor, J. A. (1999). DNA repair mechanisms for the
recognition and removal of damaged DNA bases.
Annu. Rev. Biophys. Biomol. Struct. 28:101-128.

Tainer, J. A., Thayer, M. M., and Cunningham, R. P.
(1995). DNA repair proteins. Curr. Opin. Struct.
Biol. 5:20-26.

Yang, W. (2000). Structure and function of
mismatch repair proteins. Mutat. Res.
460:245-256.

Recombination

Ortiz- Lombardia, M., Gonzalez, A., Ertja, R., Ay-
mami, J., Azorin, F., and Coll, M. (1999). Crystal
structure of a Holliday junction. Nat. Struct. Biol.
6:913-91 7.

Rafferty, J. B., Sedelnikove, S. E., Hargreaves, D.,
Artmiuk, P. J., Baker, P J., Sharpies, G. J., Mahdi, A. A.,
Lloyd, R. G., and Rice, D. W. (1996). Crystal struc-
ture of DNA recombination protein RuvA and a
model for its binding to the Holliday junction.
Science 274:415-421.

Rao, B. J., Chiu, S. K., Bazemore, L. R., Reddy, G.,
and Radding, C. M. (1995). How specific is the
first recognition step of homologous recombina-
tion? Trends Biochem. Sci. 20:109-113.

West, S. C. (1996). The RuvABC proteins and Hol-
liday junction processing in Escherichia coli. J. Bac-
teriol. 178:1237-1241.

West, S. C. (1997). Processing of recombination
intermediates by the RuvABC proteins. Annu. Rev.
Genet. 31:213-244.

West, S. C. (2003). Molecular views of recombina-
tion proteins and their control. Nat. Rev. Mol. Cell
Biol. 4:1-11.

White, M. F., Giraud-Panis, M.-J. E., Pohler, J. R. G.,
and Lilley, D. M. J. (1997). Recognition and
manipulation of branched DNA structure by
junction-resolving enzymes. /. Mol. Biol.
269:647-664.

Wuethrich, B. (1998). Why sex? Science
281:1980-1982.

Transcription and
RNA Processing

A s we have seen, the structure of DNA proposed by Watson and Crick in 1953
immediately suggested a means of replicating DNA to transfer genetic informa-
tion from one generation to the next but it did not reveal how an organism
makes use of the information stored in its genetic material.

Based on studies of the bread mold Neurospora crassa , George Beadle and Edward
Tatum proposed that a single unit of heredity, or gene, directed the production of a sin-
gle enzyme. A full demonstration of the relationship between genes and proteins came
in 1956 when Vernon Ingram showed that hemoglobin from patients with the heritable
disease sickle-cell anemia differed from normal hemoglobin by the replacement of a
single amino acid. Ingrams results indicated that genetic changes can manifest themselves
as changes in the amino acid sequence of a protein. By extension, the information contained
in the genome must specify the primary structure of each protein in an organism.

We define a gene as a DNA sequence that is transcribed. This definition includes
genes that do not encode proteins (not all transcripts are messenger RNA). The defini-
tion normally excludes regions of the genome that control transcription but are not
themselves transcribed. We will encounter some exceptions to our definition of a
gene — surprisingly, there is no definition that is entirely satisfactory.

Many prokaryotic genomes contain several thousand genes, although some simple
bacteria have only 500 to 600 genes. Most of these are “housekeeping genes” that en-
code proteins or RNA molecules that are essential for the normal activities of all living
cells. For example, the enzymes involved in the basic metabolic processes of glycolysis
and the synthesis of amino acids and DNA are encoded by such housekeeping genes, as
are transfer RNAs and ribosomal RNAs. The number of housekeeping genes in unicel-
lular eukaryotes, such as yeast and some algae, is similar to the number in complex
prokaryotes.

"This fraction (which we shall desig-
nate "messenger RNA" or M-RNA)
amounts to only about 3% of the
total RNA. . . . The property attrib-
uted to the structural messenger of
being an unstable intermediate is one
of the most specific and novel impli-
cations of this scheme. . . . This leads
to a new concept of the mechanism
of information transfer ; where the
protein synthesizing centers (ribo-
somes) play the role of non-specific
constituents which can synthesize dif-
ferent proteins , according to specific
instructions which they receive from
the genes through M-RNA. "

— Francois Jacob and Jacques
Monod, 1961

Top: A portion of the mouse transcription factor Zif268 (dark blue) bound to DNA (light blue). Side chains from three zinc-
containing domains interact with base pairs in DNA.

633

634

CHAPTER 21 Transcription and RNA Processing

) Replication

DNA ^

Transcription

RNA

Translation

Protein
a Figure 21.1

Biological information flow. The normal flow
of biological information is from DNA to
RNA to protein.

KEY CONCEPT

Before a cell can access the genetic
information stored in its DNA, the DNA
must be transcribed into RNA.

▲ Frangois Jacob (1920-). Jacob and Monod
received the Nobel Prize in Physiology or
Medicine in 1965 for their work on the ge-
netic control of enzyme synthesis.

In addition to housekeeping genes, all cells contain genes that are expressed only
in special circumstances, such as during cell division. Multicellular organisms also
contain genes that are expressed only in certain types of cells. For example, all cells in a
maple tree contain the genes for the enzymes that synthesize chlorophyll but these
genes are expressed only in cells that are exposed to light, such as cells on the surface of
a leaf. Similarly, all cells in mammals contain insulin genes, but only certain pancreatic
cells produce insulin. The total number of genes in multicellular eukaryotes ranges
from as few as 15,000 in Drosophila melanogaster to more than 50,000 in some other
animals.

In this chapter and the next, we will examine how the information stored in DNA
directs the synthesis of proteins. A general outline of this flow of information is sum-
marized in Figure 21.1. In this chapter, we describe transcription (the process where
information stored in DNA is copied into RNA thereby making it available for either
protein synthesis or other cellular functions) and RNA processing (the post-transcrip-
tional modification of RNA molecules). We also briefly examine how gene expression is
regulated by factors that affect the initiation of transcription. In Chapter 22, we will ex-
amine translation (the process where information coded in mRNA molecules directs
the synthesis of individual proteins).

One feature of the complete pathway outlined in Figure 21.1 is that it is irreversible.
In particular, the information contained in the amino acid sequence of a protein cannot
be translated back into nucleic acid. This irreversibility of information flow is known as
the “Central Dogma” of molecular biology and was predicted by Francis Crick in 1958,
many years before the mechanisms of transcription and translation were worked out
(see Section 1.1). The original version of the Central Dogma did not rule out informa-
tion flow from RNA to DNA. Such a pathway was eventually discovered in retrovirus-
infected cells; it is known as reverse transcription.

21.1 Types of RNA

Several classes of RNA molecules have been discovered. Transfer RNA (tRNA) carries
amino acids to the translation machinery. Rihosomal RNA (rRNA) makes up much of
the ribosome. A third class of RNA is messenger RNA (mRNA), whose discovery was
due largely to the work of Francois Jacob, Jacques Monod, and their collaborators at
the Pasteur Institute in Paris. In the early 1960s, these researchers showed that ribo-
somes participate in protein synthesis by translating unstable RNA molecules
(mRNA). Jacob and Monod also discovered that the sequence of an mRNA molecule is
complementary to a segment of one of the strands of DNA. A fourth class of RNA con-
sists of small RNA molecules that participate in various metabolic events, including
RNA processing. Many of these small RNA molecules have catalytic activity. Some of
these small RNAs are regulatory molecules that can bind specifically to mRNAs and
down-regulate that messenger and the protein it encodes.

A large percentage of the total RNA in a cell is ribosomal RNA, and only a small
percentage is mRNA. But if we compare the rates at which the cell synthesizes RNA
rather than the steady state levels of RNA, we see a different picture (Table 21.1). Even
though mRNA accounts for only 3% of the total RNA in Escherichia coli , the bacterium
devotes almost one-third of its capacity for RNA synthesis to the production of mRNA.
This value may increase to about 60% when the cell is growing slowly and does not need
to replace ribosomes and tRNA. The discrepancy between steady state levels of various
RNA molecules and the rates at which they are synthesized can be explained by the dif-
fering stabilities of the RNA molecules: rRNA and tRNA molecules are extremely stable,
whereas mRNA is rapidly degraded after translation. Half of all newly synthesized
mRNA is degraded by nucleases within three minutes in bacterial cells. In eukaryotes,
the average half-life of mRNA is about ten times longer. The relatively high stability of
eukaryotic mRNA results from processing events that prevent eukaryotic mRNA from
being degraded during transport from the nucleus, where transcription occurs, to the
cytoplasm, where translation occurs.

21.2 RNA Polymerase 635

Table 21.1 The RNA content of an E. coli cell

Type

Steady state level

Synthetic type capacity 0

rRNA

83%

58%

tRNA

14%

10%

mRNA

03%

32%

RNA primers* 3

<1%

Other RNA molecules 0

<1%

°Relative amount of each type of RNA being synthesized at any instant.

b RNA primers are those used in DNA replication; they are not synthesized by RNA polymerase.

c Other RNA molecules include several RNA enzymes, such as the RNA component of RNase P.

[Adapted from Bremer, H., and Dennis, P. P. (1987). Modulation of chemical composition and other parameters
of the cell by growth rate. In Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology, Vol. 2,

F. C. Neidhardt, ed. (Washington, DC: American Society for Microbiology), pp. 1527-1542.]

21.2 RNA Polymerase

About the time that mRNA was identified, researchers in several laboratories independ-
ently discovered an enzyme that catalyzes the synthesis of RNA when provided with
ATP, UTP, GTP, CTP, and a template DNA molecule. The newly discovered enzyme was
RNA polymerase. This enzyme catalyzes DNA-directed RNA synthesis, or transcription.

RNA polymerase was initially identified by its ability to catalyze polymerization of
ribonucleotides but further study of the enzyme revealed that it does much more. RNA
polymerase is the core of a larger transcription complex just as DNA polymerase is the
core of a larger replication complex (Section 20.4). This complex assembles at one end
of a gene when transcription is initiated. During initiation, the template DNA partially
unwinds and a short piece of RNA is synthesized. In the elongation phase of transcrip-
tion, RNA polymerase catalyzes the processive elongation of the RNA chain while the
DNA is continuously unwound and rewound. Finally, the transcription complex re-
sponds to specific transcription termination signals and disassembles.

Although the composition of the transcription complex varies considerably among
different organisms, all transcription complexes catalyze essentially the same types of
reactions. We introduce the general process of transcription by discussing the reactions
catalyzed by the well-characterized transcription complex in E. coli. The more compli-
cated eukaryotic transcription complexes are presented in Section 21.5.

A. RNA Polymerase Is an Oligomeric Protein

Core RNA polymerase is isolated from E. coli cells as a multimeric protein with four dif-
ferent types of subunits (Table 21.2). Five of these subunits combine with a stoichiome-
try of ct 2 /3/3'(u to form the core enzyme that participates in many of the transcription
reactions. The large /3 and /3 ' subunits make up the active site of the enzyme; the
/3' subunit contributes to DNA binding, whereas the /3 subunit contains part of the
polymerase active site. The a subunits are the scaffold for assembly of the other sub-
units and they also interact with many proteins that regulate transcription. The role of
the small co subunit is not well characterized.

The structure of RNA polymerase holoenzyme from the bacterium Thermus
aquaticus complexed with DNA is shown in Figure 21.2. The /3 and /3' subunits form a
large groove at one end. This is where DNA binds and polymerization takes place. The
groove is large enough to accommodate about 16 base pairs of double-stranded B-DNA
and is shaped like the DNA-binding sites of DNA polymerases (such as DNA poly-
merase I; Figure 20.12). The pair of a subunits is located at the “back end” of the molecule.
This region also contacts DNA when the polymerase is actively transcribing a gene.
The a) subunit is bound to the outer surface of the /3' subunit. We will see later that var-
ious transcription factors interact with RNA polymerase by binding to the a subunits.

Table 21.2 Subunits of E. coli RNA

polymerase holoenzyme

Subunit

M r

155,600

150,600

a b

70,300°

36,500

11,000

The f3 and (3 r subunits are unrelated despite the
similarity of their names.
fa This subunit is not part of the core RNA
polymerase.

The molecular weight given is for the a subunit
found in the most common form of the
holoenzyme.

636 CHAPTER 21 Transcription and RNA Processing

Figure 21.2 ►

Thermus aquaticus (taq) RNA polymerase
holoenzyme/promoter DNA closed complex.

The template strand is dark green and the
coding strand is light green; both the -10
and -35 elements are yellow. The transcrip-
tion start site is shown in red and labeled +1.
Once the open complex forms, then tran-
scription will proceed downstream, to the
right as shown by the arrows. The a and co
subunits are shown in gray; the /3 subunit is
cyan, while the (3” subunit is pink. The a
subunit is orange.

The cr subunit of the holoenzyme plays an important role in transcription initiation.
Bacteria contain several different types of cr subunits. The major form of the holoen-
zyme in E. coli contains the subunit cr 70 (M r 70,300). The cr subunits contact DNA dur-
ing transcription initiation and bind to the core enzyme in the region of the co subunit.
The overall dimensions of RNA polymerase are 10X10X16 nm. This makes it con-
siderably larger than a nucleosome but smaller than a ribosome or a replisome.

B. The Chain Elongation Reaction

RNA polymerase catalyzes chain elongation by a mechanism almost identical to that
used by DNA polymerase (Figure 20.6). Part of the growing RNA chain is base-paired to
the DNA template strand, and incoming ribonucleoside triphosphates are tested in the
active site of the polymerase for correct hydrogen bonding to the next unpaired nu-
cleotide of the template strand. When the incoming nucleotide forms correct hydrogen
bonds, RNA polymerase catalyzes a nucleotidyl- group-transfer reaction, resulting in for-
mation of a new phosphodiester linkage and the release of pyrophosphate (Figure 21.3).

Like DNA polymerase III, RNA polymerase catalyzes polymerization in the
5' — > 3' direction and is highly processive when it is bound to DNA as part of a tran-
scription complex. The overall reaction of RNA synthesis can be summarized as

RNA n - OH + NTP > RNA„ +1 - OH + PP, (21.1)

The Gibbs free energy change for this reaction is highly favorable because of the
high concentration of NTPs relative to RNA. In addition, the RNA polymerase reaction
like the DNA polymerase reaction is thermodynamically assisted by the subsequent
hydrolysis of pyrophosphate inside the cell. Thus, two phosphoanhydride linkages are
expended for every nucleotide added to the growing chain.

RNA polymerase differs from DNA polymerase in using ribonucleoside triphos-
phates (UTP, GTP, ATP, and CTP) as substrates rather than deoxyribonucleoside
triphosphates (dTTP, dGTP, dATP, and dCTP). Another difference is that the growing
RNA strand only interacts with the template strand over a short distance (see below).
The final product of transcription is single-stranded RNA, not an RNA-DNA duplex.
Transcription is much slower than DNA replication. In E. coli , the rate of transcription
ranges from 30 to 85 nucleotides per second, or less than one-tenth the rate of DNA
replication.

21.2 RNA Polymerase 637

Growing 5'RNA
RNA strand $

3'DNA
H O

©r

0= P — o 0

H O

Template strand
of DNA

◄ Figure 21.3

Reaction catalyzed by RNA polymerase. When
an incoming ribonucleoside triphosphate
correctly pairs with the next unpaired
nucleotide on the DNA template strand, RNA
polymerase catalyzes a nucleophilic attack
by the 3'-hydroxyl group of the growing RNA
strand on the a-phosphorus atom of the
incoming ribonucleoside triphosphate.

As a result, a phosphodiester forms and
pyrophosphate is released. The subsequent
hydrolysis of pyrophosphate catalyzed by
pyrophosphatase provides additional
thermodynamic driving force for the reac-
tion. (B and B' represent complementary
bases, and hydrogen bonding between bases
is indicated by a single dashed line.)

RNA polymerase catalyzes the formation of a new phosphodiester linkage only
when the incoming ribonucleoside triphosphate fits the active site of the enzyme
precisely. A precise fit requires base stacking and appropriate hydrogen bonding be-
tween the incoming ribonucleoside triphosphate and the template nucleotide.

Despite the requirement for an accurate fit, RNA polymerase does make mistakes.
The error rate of RNA synthesis is 10 -6 (one mistake for every 1 million nucleotides in-
corporated). This rate is higher than the overall error rate of DNA synthesis because, in
contrast to most DNA polymerases, RNA polymerase does not possess an exonuclease
proofreading activity. Extreme precision in DNA replication is necessary to minimize
mutations that could be passed on to progeny but accuracy in RNA synthesis is not as
crucial to survival.

638 CHAPTER 21 Transcription and RNA Processing

21.3 Transcription Initiation

The elongation reactions of RNA synthesis are preceded by a distinct initiation step in
which a transcription complex assembles at an initiation site and a short stretch of RNA
is synthesized. The regions of DNA that serve as sites of transcription initiation are
called promoters. In bacteria, several genes are often co-transcribed from a single pro-
moter; such a transcription unit is called an operon. In eukaryotic cells, each gene usu-
ally has its own promoter. There are hundreds of promoters in bacterial cells and thou-
sands in eukaryotic cells.

The frequency of transcription initiation at any given promoter is usually related to
the need for that gene’s particular product. For example, in cells that are dividing rap-
idly, the genes for ribosomal RNA are usually transcribed frequently. Every few seconds
a new transcription complex begins transcribing at the promoter. This process gives rise
to structures such as those seen in Figure 21.4 showing multiple transcription com-
plexes on one E. coli ribosomal RNA operon. Transcripts of increasing length are ar-
rayed along the genes because many RNA polymerases transcribe the genes at the same
time. In contrast, some bacterial genes are transcribed only once every two generations.
In these cases initiation may occur only once every few hours. (Outside of the labora-
tory, the average generation time of most bacteria is many hours.)

A. Genes Have a 5' — » 3' Orientation

In Section 19.2A, we introduced the convention that single-strand nucleic acid se-
quences are written from left to right in the 5' —> 3' direction. When a sequence of double-
stranded DNA is displayed, the sequence of the top strand is written 5' — > 3' and the
sequence of the bottom, antiparallel, strand is written 3' — > 5' (left to right).

Since our operational definition of a gene is a DNA sequence that is transcribed, a
gene begins at the point where transcription starts (designated +1) and ends at the
point where transcription terminates. The beginning of a gene is called the 5' end, cor-
responding to the convention for writing sequences. Moving along a gene in the
5' — > 3' direction is described as moving “downstream” and moving in the 3' — > 5'
direction is moving “upstream.” RNA polymerization proceeds in the 5' — > 3' direction.
Consequently, in accordance with the convention for writing DNA sequences, the tran-
scription start site of a gene is shown on the left of a diagram of double-stranded DNA
and the termination site is on the right. The top strand is often called the coding strand
because its sequence corresponds to the DNA version of the mRNA that encodes the
amino acid sequence of a protein. The bottom strand is called the template strand be-
cause it is the strand used as a template for RNA synthesis (Figure 21.5). Alternatively,
the top strand may be called the sense strand to indicate that translating ribosomes at-
tempting to “read” the codons in an mRNA with this sequence will make the correct
protein. Therefore the bottom strand becomes the antisense strand because an mRNA
with this sequence will not make the correct protein. Note that RNA is synthesized in

Figure 21.4 ►

Transcription of E. coli ribosomal RNA genes.

The genes are being transcribed from left to
right. The nascent rRNA product associates
with proteins and is processed by nucleolytic
cleavage before transcription is complete.

m > - ; v-

•■•Vi *.<

bush

RNA associated with protein

■

V;. ; '• -.yy-y-sy.-

\ y--; t £ - , y*-: ; J, « V V - . T ,*

•- \

Initiation site

21.3 Transcription Initiation 639

5'end 3'end

< Gene >

Transcription
start site
+1

DNA

5 , 'wv'A T CGGACCTAGGAGCC

Coding strand

.TTCCGATATACGCA
G A 1 G

C C A G A'wv'3'

◄ Figure 21.5

Orientation of a gene. The sequence of a
hypothetical gene and the RNA transcribed
from it are shown. By convention, the gene
is said to be transcribed from the 5' end to
the 3' end but the template strand of DNA
is copied from the 3' end to the 5' end.
Growth of the ribonucleotide chain
proceeds 5' — >3'.

3'-wTAGCCTGGATCCT

CGG^ Template strand ^GGT CT^5 7

t aaggctatatgcgt a

5' p p p A C C

A UUCCGAUAUACG

UAGGAG CC G

3 '

mRNA »

Direction of transcription

the 5' — > 3' direction but the template strand is copied from its 3' end to its 5' end. Also
note that the RNA product is identical in sequence to the coding strand except that
U replaces T.

B. The Transcription Complex Assembles at a Promoter

A transcription complex forms when one or more proteins bind to the promoter sequence
and also to RNA polymerase. These DNA-binding proteins direct RNA polymerase to the
promoter site. In bacteria, the cr subunit of RNA polymerase is required for promoter recog-
nition and formation of the transcription complex.

The nucleotide sequence of a promoter is one of the most important factors affect-
ing the frequency of transcription of a gene. Soon after the development of DNA-
sequencing technology, many different promoters were examined. The start sites, the
points at which transcription actually begins, were identified, and the regions upstream
of these sites were sequenced to learn whether the promoter sequences of different genes
were similar. This analysis revealed a common pattern called a consensus sequence — a
hypothetical sequence made up of the nucleotides found most often in each position.

The consensus sequence of the most common type of promoter in E. coli is shown
in Figure 21.6. This promoter is bipartite, which means that there are two separate re-
gions of sequence similarity. The first region is 10 bp upstream of the transcription start
site and is rich in A/T base pairs. The consensus sequence is TATAAT. The second part of
the promoter sequence is centered approximately 35 bp upstream of the start site. The
consensus sequence in this region is TTGACA. The average distance between the two
parts of the promoter is 17 bp.

The — 10 region is known as a TATA box, and the —35 region is simply referred to as
the -35 region. Together, the two regions define the promoter for the E. coli holoenzyme
containing cr 70 , the most common cr subunit in E. coli cells. The cr 70 -c ontaining holoen-
zyme binds specifically to sequences that resemble the consensus sequence. Other E. coli
cr subunits recognize and bind to promoters with quite different consensus sequences
(Table 21.3). Orthologous cr subunits from other prokaryotic species may recognize dif-
ferent promoter consensus sequences.

A consensus sequence is not an exact sequence but indicates the nucleotides most
commonly found at each position. Very few promoters match their consensus sequence
exactly. In some cases, the match is quite poor, with G or C found at positions normally
occupied by A or T. Such promoters are known as weak promoters and are usually asso-
ciated with genes that are transcribed infrequently. Strong promoters, such as the pro-
moters for ribosomal RNA operons, resemble the consensus sequence quite closely.
These operons are transcribed very efficiently. Observations such as these suggest that
the consensus sequence describes the most efficient promoter sequence for the RNA
polymerase holoenzyme.

KEY CONCEPT

Promoter sequences contain the
information that instructs transcription
complexes: “Initiate a transcript here.”

640

CHAPTER 21 Transcription and RNA Processing

Transcription
start site

G T G C G T G
G G C G G T G
T G A G C T G
C C C A G G C
C C C A G G C
A T C C T A C
T T T C C T C
T A A A T G C
T C C A T G T
T T A T T C C
Consensus

sequence: TTGACA TATAAT

T T G A C T
TTGACA
TTGACA
T T T A C A
T T T A C A
C T G A C G
T T G T C A
T T G A C T
C A C A C T
A T G T C A

ATTTTA CCTCTGGCGGT
TAAATA CCACT GGCGGT
ATTAAT CAT CGAACT AG
CTTTAT GCTT CCGGCT CG
CTTTAT GCTT CCGGCT CG
CTTTTT ATCGCAACTCTC
GGCCGG AATAACT CCC
CTGTAG CGGGAAGGCG
TTTCGCAT CTTTGTTATGC
CACTTT TCGCATCTTTGT

G A T A A T
G A T A C T
T T A A C T
T A T G T T
TATAAT
T A C T G T
TATAAT
T A T T A T
T A T G G T
T A T G C T

G G T T G C
G A G C A C
AG T A C G C
G T G T G G
G T G T G G
TTCTCCAT
GCGCCACC
G C A C A C C
T A T T T C
AT G G T T

T G T
T C A
A G T
ATT
ATT
CCC
C T G
C G C
T A C
T T T

ACTA
G C A G
T C A C
G T G A
G T G A
G T T T
A C A C
G C C G
C A T A
C A T A

- 35 region

- 10 region

+ 1

▲ Figure 21.6

Promoter sequences from ten bacteriophage and bacterial genes. All these promoter sequences are recognized by the a 70 subunit
cleotide sequences are aligned so that their +1, -10, and -35 regions are in register. Note the degree of sequence variation at
consensus sequence was derived from a much larger database of more than 300 well-characterized promoters.

A G G A

APr

G A C G

AP l

G T A A

trp

G C G G

lac

G C G G

/acUV5

T T T T

a ra BAD

G G A A

rrnAI

C T G A

rrnA2

A G C C

gal PI

C C A T

galP2

in E. coli. The nu-
each position. The

The promoter sequence of each gene has likely been optimized by natural selection
to fit the requirements of the cell. An inefficient promoter is ideal for a gene whose
product is not needed in large quantities whereas an efficient promoter is necessary for
producing large amounts of a gene product.

C. The cr Subunit Recognizes the Promoter

The effect of cr subunits, also called cr factors, on promoter recognition can best be ex-
plained by comparing the DNA-binding properties of core polymerase versus the
holoenzyme containing cr 70 . The core polymerase, which lacks a cr subunit, binds to
DNA nonspecifically; it has no greater affinity for promoters than for any other DNA
sequence (the association constant, IC a , is approximately 10 10 M -1 ). Once formed, this
DNA-protein complex dissociates slowly (ti/ 2 ~ 60 minutes). In contrast, the holoen-
zyme, which contains the cr 70 subunit, binds more tightly to promoter sequences
(fC a ~ 2 X 10 11 M _1 ) than the core polymerase and forms more stable complexes
(fi /2 ~ 2 to 3 hours). Although the holoenzyme binds preferentially to promoter
sequences, it also has appreciable affinity for the rest of the DNA in a cell

Table 21.3 E. coli cr subunits

Subunit Gene Genes transcribed Consensus

-35 -1 0

a 70

rpoD

Many

TTGACA

TATAAT

a 54

rpoN

Nitrogen

metabolism

None

CTGGCACNNNNNTTGCA*

<r 38

rpoS

Stationary phase

TATAAT

<r 28

flat

Flagellar synthesis
and chemotaxis

TAAA

GCCGATAA

<r 32

rpoH

Heat shock

CTTGAA

CCCATNTA 0

0-9P55

gene 55

Bacteriophage T4

None

TATAAATA

'N represents any nucleotide.

21.3 Transcription Initiation 641

(X a ~ 5 X 10 6 M -1 ). The complex formed by nonspecific binding of the holoenzyme
to DNA dissociates rapidly (ti/ 2 ~ 3 seconds). These binding parameters reveal the
functions of the cr 70 subunit. One of the roles of cr 70 is to decrease the affinity of the
core polymerase for nonpromoter sequences. Another equally important role is to in-
crease the affinity of the core polymerase for specific promoter sequences.

The association constants do not tell us how the RNA polymerase holoenzyme
finds the promoter. We might expect the holoenzyme to search for the promoter by
continuously binding and dissociating until it encounters a promoter sequence. Such
binding would be a second- order reaction, and its rate would be limited by the rate at
which the holoenzyme diffuses in three dimensions. However, promoter binding is 100
times faster than the maximum theoretical value for a diffusion-limited second-order
reaction. This remarkable rate is achieved by one-dimensional diffusion of RNA poly-
merase along the length of the DNA molecule. During the short period of time that the
enzyme is bound nonspecifically, it can scan 2000 bp in its search for a promoter se-
quence. Several other sequence-specific DNA-binding proteins, such as restriction
enzymes (Section 19.6C), locate their binding sites in a similar manner.

D. RNA Polymerase Changes Conformation

Initiation of transcription is slow, even though the holoenzyme searches for and binds
to the promoter very quickly. In fact, initiation is often the rate limiting step in tran-
scription because it requires unwinding of the DNA helix and synthesis of a short
stretch of RNA that serves as a primer for subsequent chain elongation. During DNA
replication these steps are carried out by a helicase and a primase but in transcription
these steps are carried out by the RNA polymerase holoenzyme itself. Unlike DNA poly-
merases, RNA polymerases can initiate polynucleotide synthesis on their own in the
presence of initiation factors such as cr 70 (when a DNA template and rNTPs are avail-
able as substrates).

The unwinding of DNA at the initiation site is an example of a conformational
change in which RNA polymerase (R) and the promoter (P) shift from a closed complex
(RP C ) to an open complex (RP 0 ). In the closed complex, the DNA is double-stranded.
In the open complex, 18 bp of DNA are unwound, forming a transcription bubble. For-
mation of the open complex is usually the slowest step of the initiation events.

Once the open complex forms, the template strand is positioned at the polymeriza-
tion site of the enzyme. In the next step, a phosphodiester linkage forms between two ri-
bonucleoside triphosphates that have diffused into the active site and formed hydrogen
bonds with the + 1 and +2 nucleotides of the template strand. This initiation reaction is
slower than the analogous polymerization reaction during chain elongation where one
of the substrates (the growing RNA chain) is held in place by the formation of a short
RNA-DNA helix.

Additional nucleotides are then added to the dinucleotide to create a short RNA
that is paired with the template strand. When this RNA is approximately ten nucleotides
long, the RNA polymerase holoenzyme undergoes a transition from the initiation to the
elongation mode, and the transcription complex moves away from the promoter along
the DNA template. This step is called promoter clearance. The initiation reactions can
be summarized as

^assoc

R + P < > RP C > RP 0 > (21.2)

conformational change promoter clearance

As noted earlier, the holoenzyme containing cr factor has a much greater affinity
for the promoter sequence than for any other DNA sequence. Because of this tight
binding, it resists moving away from the initiation site. However, during elongation,
the core polymerase binds nonspecifically to all DNA sequences to form a highly pro-
cessive complex. The transition from initiation to chain elongation is associated with
a conformational change in the holoenzyme that causes release of the cr subunit.
Without cr, the enzyme no longer binds specifically to the promoter and is able to
leave, or exit, the initiation site. At this time, several accessory proteins bind to the

The binding properties of RNA poly-
merase tell us that many RNA poly-
merase molecules will be located on
random stretches of DNA that may, or
may not, resemble a promoter sequence.

In transcription

complex

( 50 %)

▲ RNA polymerase distribution. Estimate of
the distribution of the approximately 5000
RNA polymerase molecules typically found
in an E. coli cell. Very few molecules are
free in the cytosol, yet only half of all RNA
polymerases are actively transcribing.

642 CHAPTER 21 Transcription and RNA Processing

◄ Figure 21.7

Initiation of transcription in E. coli.

(a) RNA polymerase holoenzyme binds nonspecifically
to DNA.

(b) The holoenzyme conducts a one-dimensional search for
a promoter.

Transcription

bubble

RNA

(d) A conformational change from the closed complex to
an open complex produces a transcription bubble at the
initiation site. A short stretch of RNA is then synthesized.

NusA

RNA

(e) The a subunit dissociates from the core enzyme, and
RNA polymerase clears the promoter. Accessory
proteins, including NusA, bind to the polymerase.

21.4 Transcription Termination 643

core polymerase to create the complete protein machine required for RNA chain
elongation. The binding of one of these accessory proteins, NusA, helps convert RNA
polymerase to the elongation form. The elongation complex is responsible for most
of the synthesis of RNA. NusA also interacts with other accessory proteins and plays a
role in termination. Transcription initiation in E. coli is summarized in Figure 21.7.

21.4 Transcription Termination

Only certain regions of DNA are specifically transcribed. Transcription complexes
assemble at promoters and, in bacteria, disassemble at the 3' end of genes at specific
sequences called termination sequences. There are two types of transcription termination
sequences. The simplest form of termination occurs at certain DNA sequences where
the elongation complex is unstable, and the transcription complex spontaneously disas-
sembles. The other type of termination requires a specific protein named rho that facili-
tates disassembly of the transcription complex, template, and mRNA.

Transcription termination often occurs near pause sites. These are regions of the
gene where the rate of elongation slows down or stops temporarily. For example, be-
cause it is more difficult to melt G/C base pairs than it is to melt A/T base pairs, a tran-
scription complex pauses when it encounters a GC-rich region.

Pausing is exaggerated at sites where the DNA sequence is palindromic, or has
dyad symmetry (Section 19.6C). When the DNA is transcribed, the newly synthesized
RNA can form a hairpin (Figure 21.8). (A three-dimensional representation of such a
structure is shown in Figure 19.21.) Formation of an RNA hairpin may destabilize the
RNA- DNA hybrid in the elongation complex by prematurely stripping off part of the
newly transcribed RNA. This partial disruption of the transcription bubble probably
causes the transcription complex to cease elongation until the hybrid re-forms. NusA
increases pausing at palindromic sites, perhaps by stabilizing the hairpin. The tran-
scription complex may pause for 10 seconds to 30 minutes, depending on the structure
of the hairpin.

Some of the strong pause sites in E. coli are termination sequences. Such termination
sites are found at the 3' end of a gene beyond the region that encodes the polypeptide
chain (for protein-encoding genes) or the complete functional RNA (for other genes).
These sites specify an RNA hairpin structure that is weakly bound to the template

◄ Figure 21.8

Formation of an RNA hairpin. The transcribed
DNA sequence contains a region of dyad
symmetry. Complementary sequences in
RNA can base-pair to form a hairpin.

S'-vw'A CCU C A C U' w ^3 /

G A

G — C
C — G
U— A

C — G RNA

A---U hairpin

G — C
G — C
A U

Dyad

symmetry

5'^ a CCTGGCTCAGGACCTT CCTGAGCACACT 3'
3'^T GGACCGAGT C C T GGAAGGACT CGT GT G A^ 5'

DNA

S'-wv'A CCUGGCUCAGGACCUUCCUGAGCACACU'

3' RNA

644 CHAPTER 21 Transcription and RNA Processing

Figure 21.9 ►

fl/zo-dependent termination of transcription in
E. coli. RNA polymerase is stalled at a
pause site where rho binds to newly synthe-
sized RNA. This binding is accompanied by
ATP hydrolysis. Rho probably wraps the nas-
cent RNA chain around itself, thereby
destabilizing the RNA-DNA hybrid and
terminating transcription.

[Adapted from Platt, T. (1986). Transcription ter-
mination and the regulation of gene expression.
Annu. Rev. Biochem. 55:339-372.]

strand by a short stretch of A/U base pairs. These are the weakest possible base pairs
(Table 19.3) and they are easily disrupted during pausing. Disruption leads to release of
RNA from the transcription complex.

The other type of bacterial termination sequences are said to be r/zo-dependent.
Rho also triggers disassembly of transcription complexes at some pause sites. It is a
hexameric protein with a potent ATPase activity and an affinity for single-stranded
RNA. Rho may also act as an RNA-DNA helicase. It binds to single-stranded RNA
that is exposed behind a paused transcription complex in a reaction coupled to hy-
drolysis of ATR Approximately 80 nucleotides of RNA wrap around the protein,
causing the transcript to dissociate from the transcription complex (Figure 21.9).
R/zo-dependent termination results from both destabilization of the RNA-DNA
hybrid and direct contact between the transcription complex and rho as rho binds
RNA. Rho can also bind to accessory proteins, such as NusA. This interaction may
cause the RNA polymerase to change conformation and dissociate from the tem-
plate DNA.

R/zo-dependent termination requires exposure of single- stranded RNA. In bacte-
ria, RNA transcribed from protein-encoding genes is typically bound by translating
ribosomes that interfere with rho binding. Single-stranded RNA only becomes ex-
posed to rho when transcription passes beyond the point where protein synthesis ter-
minates. Transcription terminates at the next available pause site. In other words, rho-
dependent termination does not occur at pause sites within the coding region but can
occur at pause sites past the translation termination codon. The net effect is to couple
transcription termination to translation. The advantages of such a coupling mechanism
are that synthesis of an mRNA coding region is not interrupted (which would prevent
protein synthesis) and that there is minimal wasteful transcription downstream of the
coding region.

21.5 Transcription in Eukaryotes 645

21.5 Transcription in Eukaryotes

The same processes carried out by a single RNA polymerase in E. coli are carried out in
eukaryotes by several similar enzymes. The activities of eukaryotic transcription com-
plexes also require many more accessory proteins than those seen in bacteria.

KEY CONCEPT

Eukaryotic transcription complexes tend
to have more factors than the analogous
bacterial complexes.

A. Eukaryotic RNA Polymerases

Three different RNA polymerases transcribe nuclear genes in eukaryotes. Other RNA
polymerases are found in mitochondria and chloroplasts. Each nuclear enzyme tran-
scribes a different class of genes (Table 21.4). RNA polymerase I transcribes genes that
encode large ribosomal RNA molecules (class I genes). RNA polymerase II transcribes
genes that encode proteins and a few that encode small RNA molecules (class II genes).
RNA polymerase III transcribes genes that encode a number of small RNA molecules,
including tRNA and 5S rRNA (class III genes). (Some of the RNA molecules listed in
the table are discussed in subsequent sections.)

The mitochondrial version of RNA polymerase is a monomeric enzyme encoded
by the nuclear genome. It is substantially similar in amino acid sequence to the RNA
polymerases of T3 and T7 bacteriophages. This similarity suggests that these enzymes
share a common ancestor. It is likely that the gene for mitochondrial RNA polymerase
was transferred to the nucleus from the primitive mitochondrial genome.

Chloroplast genomes often contain genes that encode their own RNA poly-
merase. The genes encoding the chloroplast RNA polymerase are similar in sequence
to those of RNA polymerase in cyanobacteria. This is further evidence that chloro-
plasts, like mitochondria, originated from bacterial endosymbionts in ancestral
eukaryotic cells.

The three nuclear RNA polymerases are complex multisubunit enzymes. They dif-
fer in subunit composition, although they share several small polypeptides in common.
The exact number of subunits in each polymerase varies among organisms but there are
always 2 large subunits and 7 to 12 smaller ones (Figure 21.10). RNA polymerase II
transcribes all protein-coding genes as well as some genes that encode small RNA mole-
cules. The protein- coding RNA synthesized by this enzyme was originally called hetero-
geneous nuclear RNA (hnRNA) but it is now more commonly referred to as mRNA
precursor, or pre-mRNA. The processing of this precursor into mature mRNA is de-
scribed in Section 21.9.

About 40,000 molecules of RNA polymerase II are found in large eukaryotic cells;
the activity of this enzyme accounts for roughly 20% to 40% of all cellular RNA synthe-
sis. The two largest subunits of each nuclear eukaryotic RNA polymerase are similar in
sequence to the (3 and (3 ' subunits of E. coli RNA polymerase indicating that they
share a common ancestor. Like their prokaryotic counterparts, the core eukaryotic RNA

Table 21.4 Eukaryotic RNA polymerases

Polymerase

Location

Copies
per cell

Products

Polymerase
activity of cell

RNA polymerase 1

Nucleolus

40,000

35-47S pre-rRNA

50%-70%

RNA polymerase II

Nucleoplasm

40,000

mRNA precursors

U1, U2, U4, and U5 snRNA

20%-40%

RNA polymerase III

Nucleoplasm

20,000

5S rRNA
tRNA
U6 snRNA
7S RNA
Other small
RNA molecules

10%

Mitochondrial
RNA polymerase

Mitochondrion

Products of all mitochondrial
genes

<1%

Chloroplast RNA
polymerase

Chloroplast

Products of all chloroplast
genes

<1%

646 CHAPTER 21 Transcription and RNA Processing

Figure 21.10 ►

RNA polymerase II from the yeast
Saccharomyces cerevisiae. The large subunit
colored purple (Rpb2) is the homolog of the
/ 3 subunit of the prokaryotic enzyme shown
in Figure 21.2. [PDB 1EN0].

Rpb2

polymerases do not bind on their own to promoters. RNA polymerase II requires
five different biochemical activities, or factors, to form a basal transcription complex
capable of initiating transcription on a minimal eukaryotic promoter (Figure 21.11).
These general transcription factors (GTFs) are: TFIIB, TFIID, TFIIE, TFIIF and TFIIH
(Table 21.5).

Many class II genes contain an A/T-rich region, also called a TATA box, that is func-
tionally similar to the prokaryotic TATA box discussed above (recall that A/T-rich
regions are more easily unwound to create an open complex, especially if the DNA is
negatively supercoiled (Section 19.3)). This eukaryotic A/T-rich region is located 19 to
27 bp upstream of the transcription start site and serves to recruit RNA polymerase II
to the DNA during assembly of the initiation complex.

The general transcription factor TFIID is a multisubunit factor and one of its sub-
units, TATA-binding protein (TBP), binds to the region containing the TATA box. The
structure of TBP from the plant Arabidopsis thaliana is shown in Figure 21.12. TBP
forms a saddle-shaped molecular clamp that almost surrounds the DNA at the TATA
box. The main contacts between TBP and DNA are due to interactions between acidic
amino acid side chains in (3 strands and the edges of base pairs in the minor groove.
When TBP binds to DNA, the promoter DNA is bent so that it no longer resembles the
standard B-DNA conformation. This is an unusual interaction for DNA-binding pro-
teins. The TBP subunit of TFIID is also required to initiate transcription of class I and
class III genes by RNA polymerases I and III, respectively.

The eukaryotic RNA polymerase II subunit homologous to the prokaryotic RNA
polymerase /3' subunit has an unusual carboxy- terminal domain (CTD) or “tail” that

Figure 21.11 ►

A generic eukaryotic promoter showing the
basal or “core” promoter elements. The TATA
box is described in the text. The BRE is the
TFIIB recognition element, while Inr stands
for the initiator element. The DPE is the
downstream promoter element. The names of
the factors that bind to each site are shown
above the promoter, and the consensus
recognition sequences for each site are shown
below the schematic promoter fragment.

5' TFIIB TBP

-37 -32-31 -26

TFIID TFIID 3'

-2 +4 +28 +32

21.5 Transcription in Eukaryotes 647

Table 21.5 Some representative RNA polymerase II transcription factors

Factor

Characteristics

TFIIA

Binds to TFIID; can interact with TFIID in the absence of DNA

TFIIB

Interacts with RNA polymerase II

TFIID

RNA polymerase II initiation factor

TBP

TATA-binding protein; subunit of TFIID

TAFs

TBP-associated factors; many subunits

TFIIE

Interacts with RNA polymerase II

TFIIH

Required for initiation; helicase activity; couples transcription to DNA repair

TFIIS 0

Binds to RNA polymerase II; elongation factor

TFIIF

Binds to RNA polymerase II; two subunits — RAP30 and RAP74

SP1

Binds to GC-rich sequence

CTF b

Family of different proteins that recognize the core sequence CCAAT

°Also known as si I or RAP 3 8
fa Also known as NP1 .

consists of multiple repeats of the amino acid heptamer PTSPSYS. The Ser and Thr
residues in the tail are phosphorylation targets for nuclear protein kinases. RNA poly-
merase II molecules with a hyperphosphorylated CTD are typically transcriptionally
active, or engaged, while the cellular pol II with hypophosphorylated CTDs are usually
quiescent.

Although it has proven possible to purify RNA polymerase II and each GTF and
use them to reconstitute accurate transcription initiation in vitro , these basal transcrip-
tion complexes are not competent to recognize the many different types of trans-acting
factors and ds-acting sequences that are known to play important roles in vivo. Search-
ing for cellular constituents that could respond to transcriptional activators in vitro led
to the discovery of a large preformed RNA pol II holoenzyme that contains not only the
five GTFs but also many other polypeptides that mediate interactions between pol II
and sequence-specific DNA-binding proteins. This eukaryotic holoenzyme is analogous
to the core + cr holoenzyme in E. coli.

▲ Figure 21.12

Arabidopsis thaliana TATA-binding protein
(TBP) bound to DNA. TBP (blue) is bound to
a double-stranded DNA fragment with a
sequence corresponding to a TATA box
(5'-TATAAAG-3') DNA is shown as a wire-
frame model. Note that the (3 sheet of TBP
lies in the minor groove of the DNA fragment.
[PDB 1V0L].

B. Eukaryotic Transcription Factors

TFIIA and TFIIB are essential components of the RNA polymerase II holoenzyme complex.
Neither TFIIA nor TFIIB can bind to DNA in the absence of TFIID. TFIIF (also known
as Factor 5 or RAP30/74) binds to RNA polymerase II during initiation (Figure 21.13).
TFIIF plays no direct role in recognizing the promoter but it is analogous to bacterial cr
factors in two ways: it decreases the affinity of RNA polymerase II for nonpromoter

TBP (subunit
of TFIID)

TFIID

TFIIB

TFIIA

RNA
polymerase II

P 30/74

◄ Figure 21.13

RNA polymerase II holoenzyme complex bound
to a promoter. This model shows various tran-
scription factors bound to RNA polymerase II
at a promoter. The transcription factors are
often larger and more complex than those
shown in this diagram.

648

CHAPTER 21 Transcription and RNA Processing

DNA, and it helps form the open complex. TFIIH, TFIIE, and other, less well- characterized
factors, are also part of the transcription initiation complex.

Once the initiation complex assembles at the site of the promoter, the next
steps are similar to those in bacteria. An open complex is formed, a short stretch of
RNA is synthesized, and the transcription complex clears the promoter. Most tran-
scription factors dissociate from DNA and RNA polymerase II once elongation be-
gins. However, TFIIF may remain bound and a specific elongation factor, TFIIS
(also called sll or RAP38), associates with the transcribing polymerase. TFIIS may
play a role in pausing and transcription termination that is similar to the role of
NusA in bacteria.

With the exception of TBP, the transcription factors that interact with the other
two eukaryotic RNA polymerases are not the same as those required by RNA
polymerase II.

C. The Role of Chromatin in Eukaryotic Transcription

As described in Chapter 19, the eukaryotic genome is packaged using small, ubiqui-
tous building blocks, called nucleosomes, that contain an octamer of the four core
histone proteins. It is estimated that approximately 35% of the mammalian genome is
transcribed into protein-coding genes (including the introns) and so most of a cell’s
DNA is relatively inert. But even within that 35%, which contains about 20,000
protein-coding genes, the majority of the sequences are quiescent. In any single cell,
the primary determinant of whether a gene is competent to be transcribed resides in
the state of its chromatin. This status is modulated by two mechanisms. The first in-
volves implementing or removing post-translational modifications on the flexible
amino-terminal arms of the four core histones (Section 19.5B). Specific Lys residues
are targeted for methylation or acetylation, specific Arg residues may also be methy-
lated, while Ser and Thr side chains can be phosphorylated. Different modifications
serve as signals to recruit either activators or repressors to the chromatin. The second
mechanism for specifying the transcriptional status of a eukaryotic gene involves nu-
cleosome positioning and remodeling.

Nontranscribed genes are relatively inaccessible in the nucleus while transcribed
genes are relatively accessible to transcription factors, pol II holoenzyme, and other nu-
clear proteins. How does a gene move between these two conflicting states? The answer
lies with large multiprotein complexes that use the energy from hydrolyzing ATP to
physically remodel a gene’s nucleosomes and allow proteins to have access to the DNA.
Some of the remodeling complexes actually contain histone-modifying enzymes like
histone acetylase (HAT) or histone deacetylase (HDAC).

21.6 Transcription of Genes Is Regulated

As noted at the beginning of this chapter, many genes are expressed in every cell. The
expression of these housekeeping genes is said to be constitutive. In general, such genes
have strong promoters and are transcribed efficiently and continuously. Genes whose
products are required at low levels usually have weak promoters and are transcribed in-
frequently. In addition to constitutively expressed genes, cells contain genes that are ex-
pressed at high levels in some circumstances and not at all in others. Such genes are said
to be regulated.

Regulation of gene expression can occur at any point in the flow of biological infor-
mation but occurs most often at the level of transcription. Various mechanisms have
evolved that allow cells to program gene expression during differentiation and develop-
ment and to respond to environmental stimuli.

The initiation of transcription of regulated genes is controlled by regulatory pro-
teins that bind to specific DNA sequences. Transcriptional regulation can be negative or
positive. Transcription of a negatively regulated gene is prevented by a regulatory pro-
tein called a repressor. A negatively regulated gene can be transcribed only in the ab-
sence of an active repressor. Transcription of a positively regulated gene can be activated

21.6 Transcription of Genes Is Regulated

649

by a regulatory protein called an activator. A positively regulated gene is transcribed
poorly or not at all in the absence of the activator.

Repressors and activators are often allosteric proteins whose function is modified
by ligand binding. In general, a ligand alters the conformation of the protein and
affects its ability to bind to specific DNA sequences. For example, some repressors con-
trol the synthesis of enzymes for a catabolic pathway. In the absence of substrate for
these enzymes, the genes are repressed. When substrate is present, it binds to the re-
pressor, causing the repressor to dissociate from the DNA and allowing the genes to be
transcribed. Ligands that bind to and inactivate repressors are called inducers because
they induce transcription of the genes controlled by the repressors. In contrast, some
repressors that control the synthesis of enzymes for a biosynthetic pathway bind to
DNA only when associated with a ligand. The ligand is often the end product of the
biosynthetic pathway. This regulatory mechanism ensures that the genes in the path-
way are turned off as product of the pathway accumulates. Ligands that bind to and
activate repressors are called corepressors. The DNA-binding activity of allosteric activators
can also be affected in two ways by ligand binding. Four general strategies for regulat-
ing transcription are illustrated in Figure 21.14. Examples of all four strategies have
been identified.

Few regulatory systems are as simple as those described above. For example, the
transcription of many genes is regulated by a combination of repressors and activa-
tors or by multiple activators. Elaborate mechanisms for regulating transcription

KEY CONCEPT

Cells don’t synthesize a specific protein
until it is required (e.g., the lac operon
is not transcribed until the intracellular
concentration of lactose inactivates the
lac repressors).

Activator

RNA polymerase

(a) An activator with bound ligand
stimulates transcription.

◄ Figure 21.14

Strategies for regulating transcription
initiation by regulatory proteins.

Activator

(b) An activator stimulates

transcription. In the presence of
ligand, the activator is inhibited.

( v

(c) A repressor prevents transcription.
Binding of ligand (inducer) to the
repressor inactivates the repressor
and allows transcription.

(d) In the absence of ligand, the
repressor does not bind to DNA.
Repression occurs only when
ligand (corepressor) is present.

650

CHAPTER 21 Transcription and RNA Processing

have evolved to meet the specific requirements of individual organisms. A greater
range of cellular responses is possible when transcription is regulated by a host of
mechanisms acting together. By examining how the transcription of a few particular
genes is controlled, we can begin to understand how positive and negative mecha-
nisms can be combined to produce the remarkably sensitive regulation seen in
bacterial cells.

21.7 The lac Operon, an Example of Negative
and Positive Regulation

Some bacteria obtain the carbon they need for growth by metabolizing five- or six-carbon
sugars via glycolysis. For example, E. coli preferentially uses glucose as a carbon source
but can also use other sugars, including /3-galactosides such as lactose. The enzymes re-
quired for /3-galactoside uptake and catabolism are not synthesized unless a /3-galactoside
substrate is available. Even in the presence of their substrate, these enzymes are synthe-
sized in limited amounts when the preferred carbon source (glucose) is also present.
Synthesis of the enzymes required for /3-galactoside utilization is regulated at the level
of transcription initiation by a repressor and an activator.

The uptake and catabolism of /3-galactosides requires three proteins. The product
of the lacY gene is lactose permease, a symport transporter that is responsible for the
uptake of /3-galactosides. Most /3-galactosides are subsequently hydrolyzed to metabo-
lizable hexoses by the activity of /3-galactosidase, a large enzyme with four identical sub-
units encoded by the lacZ gene. /3-Galacto sides that cannot be hydrolyzed are acetylated
by the activity of thiogalactoside transacetylase, the product of the lacA gene. Acetyla-
tion helps to eliminate toxic compounds from the cell.

The three genes — lacZ , lacY , and lacA — form an operon that is transcribed from a
single promoter to produce a large mRNA molecule containing three separate protein-
coding regions. In this case, we refer to a protein-coding region as a gene, a definition
that differs from our standard use of the term. The arrangement of genes with related
functions in an operon is efficient because the concentrations of a set of proteins can be
controlled by transcribing from a single promoter. Operons composed of protein- coding
genes are common in E. coli and other prokaryotes but were thought to be extremely
rare in eukaryotes. We now realize that operons are also quite common in the model or-
ganism C. elegans , a nematode or round worm, and are likely widespread in this large
phylum. Operons are also common in mitochondrial and chloroplast genomes.

A. lac Repressor Blocks Transcription

Expression of the three genes of the lac operon is controlled by a regulatory protein
called lac repressor, a tetramer of identical subunits. The repressor is encoded by a
fourth gene, lacl , which is located just upstream of the lac operon but is transcribed
from a separate promoter (Figure 21.15).

lac repressor binds simultaneously to two sites near the promoter of the lac
operon. Repressor-binding sites are called operators. One operator (O x ) is adjacent to
the promoter, and the other (0 2 ) is within the coding region of lacZ. When bound to
both operators, the repressor causes the DNA to form a stable loop that can be seen

Figure 21.15 ►

Organization of the genes that encode proteins
required to metabolize lactose. The coding re-
gions for three proteins — LacZ, LacY, and
LacA — constitute the lac operon and are co-
transcribed from a single promoter (P /ac ).

The gene that encodes lac repressor, lad, is
located upstream of the lac operon and has
its own promoter, P; lac repressor binds to
the operators Oi and 0 2 near P/ac! t denotes
the transcription termination sequence.

lac operon

lacl lacZ lacY lacA

21.7 The lac Operon, an Example of Negative and Positive Regulation 651

◄ Figure 21.16

Electron micrographs of DNA loops. These
loops were formed by mixing lac repressor
with a fragment of DNA bearing two syn-
thetic lac repressor-binding sites. One bind-
ing site is located at one end of the DNA
fragment, and the other is 535 bp away.
DNA loops 535 bp in length form when the
tetrameric repressor binds simultaneously to
the two sites.

B. The Structure of lac Repressor

The role of lac repressor in regulating expression of the lac operon has been
known since the 1960s. However, the structure of this important protein
was solved only in the 1990s after the development of new techniques for
determining the structure of large molecules. The structure of part of the
lac repressor bound to one operator sequence is shown in Figure 21.19. The
complete protein contains four identical subunits arranged as two pairs,
and each pair of subunits binds to a different operator sequence. Inside
the cell these two fragments of DNA are part of a single DNA molecule —
and repressor binding forms a loop of DNA at the 5' end of the lac operon.

▲ Figure 21.17

Binding of lac repressor to the lac operon. The

tetrameric lac repressor interacts simultane-
ously with two sites near the promoter of the
lac operon. As a result, a loop of DNA forms.
RNA polymerase can still bind to the pro-
moter in the presence of the lac
repressor-DNA complex.

in electron micrographs of the complex formed between lac repressor and DNA
(Figure 21.16). The interaction of lac repressor with the operator sequences may
block transcription by preventing the binding of RNA polymerase to the lac pro-
moter. However, it is now known that, in some cases, both lac repressor and RNA
polymerase can bind to the promoter at the same time. Thus, the repressor may also
block transcription initiation by preventing formation of the open complex and
promoter clearance. A schematic diagram of lac repressor bound to DNA in the pres-
ence of RNA polymerase is shown in Figure 21.17. The diagram illustrates the rela-
tionship between the operators and the promoter and the DNA loop that forms
when the repressor binds to DNA.

The repressor locates an operator by binding nonspecifically to DNA and search-
ing by sliding or hopping in one dimension. The non-specific equilibrium constant is
about 10 6 M -1 — comparable to that of RNA polymerase (Section 21.3C). (Recall from
Section 21.3C that RNA polymerase also uses this kind of searching mechanism.) The
equilibrium association constant for the specific binding of lac repressor to Oj_ in
vitro is very high (K a ~ 10 13 M -1 ). As a result, the repressor blocks tran-
scription very effectively, (lac repressor binds to the 0 2 site with lower
affinity.) A bacterial cell contains only about ten molecules of lac repressor
but the repressor searches for and finds an operator so rapidly that when a
repressor dissociates spontaneously from the operator, another occupies
the site within a very short time. However, during this brief interval, one
transcript of the operon can be made since RNA polymerase is poised at
the promoter. This low level of transcription, called escape synthesis, en-
sures that small amounts of lactose permease and /3-galactosidase are pres-
ent in the cell.

In the absence of lactose, lac repressor blocks expression of the lac
operon, but when /3-galactosides are available as potential carbon sources,
the genes are transcribed. Several /3-galactosides can act as inducers. If
lactose is the available carbon source, the inducer is allolactose, which is
produced from lactose by the action of /3-galactosidase (Figure 21.18).
Allolactose binds tightly to lac repressor and causes a conformational
change that reduces the affinity of the repressor for the operators
(K a ~ 10 10 M -1 ). In the presence of the inducer, lac repressor dissociates
from the DNA, allowing RNA polymerase to initiate transcription. (Note
that because of escape synthesis, lactose can be taken up and converted to
allolactose even when the operon is repressed.)

At any given time, one molecule of
repressor is bound to the operator and
nine molecules are bound non-specifically
to DNA.

RNA polymerase

652

CHAPTER 21 Transcription and RNA Processing

Lactose

(/3-D-Galactopyranosyl-(1 -> 4)-/3-D-glucopyranose)

/3-Galactosidase

Allolactose

(/3-D-Galactopyranosyl-(1 6)-/3-D-glucopyranose)
▲ Figure 21.18

Formation of allolactose from lactose, cat-
alyzed by /S-galactosidase. This is a minor or
side reaction. The main enzymatic activity
of /3-galactosidase is to cleave disaccharides
into monomers that can be converted into
substrates for glycolysis.

The subunits are joined together at a hinge region. The X-ray crystallo-
graphic structure reveals that the two pairs of subunits are stacked on top of
one another (Figure 21.17) and not extended away from the hinge region as
was expected. This makes a more compact protein that is less symmetric
than many other tetrameric proteins.

Each subunit contains a helix- turn -helix motif at the ends farthest from
the hinge region. When bound to DNA, one of the a helices lies in the major
groove where amino acid side chains interact directly with the specific base
pairs of the operator sequence. The two helices from each pair of subunits
are positioned about one turn of DNA apart (about 10 bp), and each one in-
teracts with half of the operator sequence. This binding strategy is similar to
that of restriction endonuclease EcoRl (Section 19.6C).

In the absence of DNA the distal regions of the lac repressor subunits
are disordered (Section 4.7D). This is one reason why it took such a long
time to work out the structure. The structure of the helix-turn-helix motif
can only be seen when the protein is bound to DNA. There are now many
examples of such interactions in which the stable structure of the protein is
significantly altered by ligand binding. In the presence of inducers, such as
allolactose or IPTG, the repressor adopts a slightly different conformation
and can no longer bind to the DNA operators.

C. cAMP Regulatory Protein Activates Transcription

Transcription of the lac operon in E. coli depends not only on the presence
of /3-galactosides but also on the concentration of glucose in the external
medium. The lac operon is transcribed maximally when /3-galactosides,
such as lactose, are the only carbon source; transcription is reduced 50-fold
when glucose is also present. The decreased rate of transcription of operons
when glucose is present is termed catabolite repression.

Catabolite repression is a feature of many operons encoding meta-
bolic enzymes. These operons characteristically have weak promoters
from which transcription is initiated inefficiently in the presence of glu-
cose. In the absence of glucose, however, the rate of transcription initia-
tion increases dramatically due to an activator that converts the relatively
weak promoter to a stronger one. No repressor is involved, despite the

Figure 21.19 ►

Structure of E. coli lac repressor. This figure
shows a dimer of lac repressor subunits
bound to DNA. Lac repressor is a tetramer
in vivo, containing two DNA-binding sites.

(a) End-on view of the DNA molecule.

(b) Side view showing the lac repressor a
helix in the major groove. [PDB 1EFA].

21.7 The lac Operon, an Example of Negative and Positive Regulation 653

use of the term catabolite repression. In fact, this is a well-studied
example of an activation mechanism.

The activator is cyclic AMP regulatory (or receptor) protein
(CRP), also known as catabolite activator protein (CAP). CRP is a
dimeric protein whose activity is modulated by cyclic AMP. In the ab-
sence of cAMP, CRP has low affinity for DNA but when cAMP is
present it binds to CRP and converts it to a sequence-specific
DNA-binding protein. The CRP- cAMP complex interacts with specific
DNA sequences near the promoters of more than 30 genes including
the lac operon. Because the genome contains many more binding
sites for CRP-cAMP than for lac repressor, it is not surprising that
there are at least 1000 molecules of CRP per cell compared to only
about 10 molecules of lac repressor. The CRP-cAMP binding sites are
often just upstream of the —35 regions of the promoters they acti-
vate. While bound to DNA, CRP-cAMP can contact RNA polymerase
at the promoter site, leading to increased rates of transcription initia-
tion (Figure 21.20). Most of the protein-protein interactions are be-
tween bound CRP-cAMP and the a subunits of RNA polymerase.
This is typical of most interactions between activators and RNA
polymerase. (There are many different transcriptional activators in
bacterial cells.) The net effect of CRP-cAMP is to increase the pro-
duction of enzymes that can use substrates other than glucose. In the
case of the lac operon, activation by CRP-cAMP occurs only when
/3-galactosides are available. At other times, transcription of the
operon is repressed.

The concentration of cAMP inside an E. coli cell is controlled
by the concentration of glucose outside the cell. When glucose is avail-
able, it is imported into the cell and phosphorylated by a complex of
transport proteins collectively known as the phosphoenolpyruvate-
dependent sugar phosphotransferase system. When glucose is not avail-
able, one of the glucose transport enzymes, enzyme III, catalyzes the
transfer of a phosphoryl group, ultimately derived from phospho-
enolpyruvate, to adenylate cyclase, leading to its activation (Figure 21.21).

(a) CRP-cAMP binds to a site near the
promoter.

DNA

RNA

polymerase

holoenzyme

Promoter

CRP-cAMP

(b) RNA polymerase holoenzyme binds to
the promoter and also contacts the bound
activator, which increases the rate of
transcription initiation.

CRP-cAMP

Promoter

▲ Figure 21.20

Activation of transcription initiation at the lac
promoter by CRP-cAMP.

Plasma

membrane

Cytosol

cAMP + PP;

HPr

r Pyruvate
Phosphoenolpyruvate

CRP-cAMP

◄ Figure 21.21

cAMP production. In the absence of glucose,
enzyme III (EMI) transfers a phosphoryl
group, originating from phosphoenolpyru-
vate, to membrane-bound adenylate cyclase.
Phosphorylated adenylate cyclase catalyzes
the conversion of ATP to cAMP. cAMP binds
to CRP, and CRP-cAMP activates the tran-
scription of a number of genes encoding
enzymes that compensate for the lack of
glucose as a carbon source.

654 CHAPTER 21 Transcription and RNA Processing

CAMP

binding

Adenylate cyclase (also know as adenylyl cyclase) then catalyzes the conversion of
ATP to cAMP thereby increasing the levels of cAMP in the cell. As molecules of
cAMP are produced, they bind to CRP stimulating transcription initiation at pro-
moters that respond to catabolite repression. Similar mechanisms for responding
to external stimuli operate in eukaryotes where molecules such as cAMP act as sec-
ond messengers (Section 9.12B).

Each subunit of the CRP dimer contains a helix-turn-helix DNA binding
motif. In the presence of cAMP, two helices — one from each monomer — fit into
adjacent sections of the major groove of DNA and contact the nucleotides of the
CRP-cAMP binding site. This is the same general binding strategy used by lac re-
pressor and EcoRl. In the absence of cAMP, the conformation of CRP changes so
that the two a helices can no longer bind to the major groove (Figure 21.22).
When CRP-cAMP is bound to the activator sequence, the DNA is bent slightly to
conform to the surface of the protein (Figure 21.23).

cAMP

a helix

cAMP

helix

▲ Figure 21.22

Conformational changes in CRP caused by
cAMP binding. Each monomer of the CRP
dimer contains a helix-turn-helix motif. In
the absence of cAMP, the a helices cannot
fit into adjacent sections of the major groove
of DNA and cannot recognize the CRP-cAMP
binding site. When cAMP binds to CRP, the
two a helices assume the proper conforma-
tion for binding to DNA.

21.8 Post-transcriptional Modification
of RNA

In many cases, RNA transcripts must be extensively altered before they can adopt
their mature structures and functions. These alterations fall into three general cat-
egories: (1) removal of nucleotides from primary RNA transcripts; (2) addition of
nucleotides not encoded by the corresponding genes; and (3) covalent modifica-
tion of certain bases. The reactions that transform a primary RNA transcript into a
mature RNA molecule are referred to collectively as RNA processing. RNA process-
ing is crucial for the function of most RNA molecules and is an integral part of
gene expression.

A. Transfer RNA Processing

Mature tRNA molecules are generated in both eukaryotes and prokaryotes by pro-
cessing primary transcripts. In prokaryotes, the primary transcript often contains
several tRNA precursors. These precursors are cleaved from the large primary
transcripts and trimmed to their mature lengths by ribonucleases, or RNases.
Figure 21.24 summarizes the processing of prokaryotic tRNA precursors.

The endonuclease RNase P catalyzes the initial cleavage of most tRNA pri-
mary transcripts. The enzyme cleaves the transcript on the 5' side of each tRNA
sequence, releasing monomeric tRNA precursors with mature 5' ends. Diges-
tion with RNase P in vivo is rapid and occurs while the transcript is still being
synthesized.

Figure 21.23 ►

Structure of a complex between CRP-cAMP

and DNA. Both subunits contain a molecule
of cAMP bound at the allosteric site. Each
subunit has an a helix positioned in the
major groove of DNA at the CRP-cAMP
binding site. Note that binding induces a
slight bend in the DNA. [PDB 1CGP].

21.9 Eukaryotic mRNA Processing 655

RNase P was one of the first specific ribonucleases studied in detail and much is
known about its structure. The enzyme is actually a ribonucleoprotein. In E. coli, it is
composed of a 377-nucleotide RNA molecule (M r 130,000) and a small polypeptide
(M r 18,000). In the absence of protein the RNA component is catalytically active in vitro
(under certain conditions). It was one of the first RNA molecules shown to have
enzymatic activity and is an example of the fourth class of RNA molecules described in
Section 21.1. The protein component of RNase P helps maintain the three-dimensional
structure of the RNA. Sidney Altman was awarded the Nobel Prize in 1989 for showing
that the RNA component of RNase P had catalytic activity.

Other endonucleases cleave tRNA precursors near their 3' ends. Subsequent pro-
cessing of the 3' end of a tRNA precursor requires the activity of an exonuclease, such as
RNase D. This enzyme catalyzes the sequential removal of nucleotides from the 3' end
of a monomeric tRNA precursor until the 3 ' end of the tRNA is reached.

All mature prokaryotic and eukaryotic tRNA molecules must contain the sequence
CCA as the final three nucleotides at their 3' ends. In some cases, these nucleotides are
added post-transcriptionally after all other types of processing at the 3' end have been
completed. The addition of these three nucleotides is catalyzed by tRNA nucleotidyl-
transferase and is one of the few examples of the addition of nucleotides that are not
encoded by a gene.

Processing of tRNA precursors also involves covalently modifying some of the nu-
cleotide bases. Mature tRNA molecules exhibit a greater diversity of covalent modifica-
tions than any other class of RNA molecule. Typically 26 to 30 of the approximately 80
nucleotides in a tRNA molecule are covalently modified. Each type of covalent modifi-
cation usually occurs in only one location on each molecule. Some examples of the sites
of modification of nucleotides are shown in Figure 21.25.

B. Ribosomal RNA Processing

Ribosomal RNA molecules in all organisms are produced as large primary transcripts
that require subsequent processing, including methylation and cleavage by endonucle-
ases, before the mature molecules can adopt their active forms. This processing of ribo-
somal RNA is coupled to ribosome assembly.

The primary transcripts of prokaryotic rRNA molecules are about 3 OS in size and
contain one copy each of the 16S, 23S, and 5S rRNAs. The transcripts also contain inter-
spersed tRNA precursors. (Note that S is the symbol for the Svedberg unit, a measure of
the rate at which particles move in the gravitational field established in an ultracen-
trifuge. Large S values are associated with large masses. The relationship between S and
mass is not linear; therefore, S values are not additive.) Since the three rRNAs are de-
rived from a single transcript, this processing ensures that there are equimolar amounts
of each of the mature ribosomal RNAs.

The 5' and 3' ends of each mature rRNA molecule are usually found in base-paired
regions in the primary transcript. In prokaryotes, the endonuclease RNase III binds to
these regions and cleaves the precursor near the ends of the 16S and 23S rRNAs. Follow-
ing the initial cleavage, the ends of the rRNA molecules are trimmed by the actions of
specific endonucleases (Figure 21.26).

Eukaryotic ribosomal RNAs are also produced by processing a larger precursor.
The primary transcripts are between 35S and 47S in size and contain a copy of each of
three eukaryotic rRNA species: 18S, 5.8S, and 28S. (The fourth eukaryotic rRNA, 5S
rRNA, is transcribed as a monomer by RNA polymerase III and is processed separately.)
The primary transcripts are synthesized in the region of the nucleus called the nucleolus,
where initial processing occurs. Each rRNA precursor partially folds up and binds to
some of its ribosomal protein partners before the processing cleavages take place.

(a) RNase P and other endonucleases
cleave the primary transcript.

RNase P

RNase D

2 CTP + ATP

▲ Figure 21.24

Summary of prokaryotic tRNA processing.

21.9 Eukaryotic mRNA Processing

KEY CONCEPT

The processing of mRNA precursors is one of the biochemical features that distin- Unmodified mRNAs are inherently

guishes prokaryotes from eukaryotes. In prokaryotes, the primary mRNA tran- unstable in a cell and would be rapidly

scripts are translated directly, often initiating translation before transcription is degraded by ribonucleases.

656

CHAPTER 21 Transcription and RNA Processing

A/ 6 -Methyladenylate

(m 6 -A)

Dihydrouridylate

(D)

▲ Figure 21.25

Examples of common covalent modifications
found in tRNA molecules (the modifications are
shown in blue).

Figure 21.26 ►

Endonucleolytic cleavage of ribosomal RNA
precursors in E. coli. The primary transcript
contains a copy of each of the three riboso-
mal RNAs and may also contain several
tRNA precursors. The large rRNA precursors
are cleaved from the large primary transcript
by the action of RNase III. The ends of the
16S, 23S, and 5S rRNAs are trimmed by
the action of endonucleases M16, M23, and
M5, respectively. (Slash marks indicate that
portions of the rRNA primary transcript have
been deleted for clarity.)

A/ 6 -lsopentenyladenylate

(i 6 -A)

Inosinate

(I)

7-Methylguanylate

(m 7 G)

Pseudouridylate

(ribose at C-5)

Uridylate
5-oxyacetic acid
(cmo 5 -U)

3-Methylcytidylate

(m 3 C)

5-Methylcytidylate

(m 5 C)

2'-0-Methylated

nucleotide

(Nm)

21.9 Eukaryotic mRNA Processing

657

complete. In eukaryotes, on the other hand, transcription occurs in the nucleus,
and translation takes place in the cytoplasm. This compartmentalization of func-
tions in eukaryotic cells allows nuclear processing of mRNA precursors without
disrupting translation.

Mature eukaryotic mRNA molecules are often derived from much larger primary
transcripts. Subsequent processing of these primary transcripts includes some of the
same steps that we saw in the previous section, namely: cleavage of a precursor, addition
of terminal nucleotides, and covalent modification of nucleotides. Often, specific nu-
cleotides (called intervening sequences, or introns) from the middle of an mRNA pri-
mary transcript are actually excised, or removed, and the resulting fragments are ligated
together to produce the mature mRNA. This step, called splicing, is common in most
eukaryotic species. Splicing also occurs during the processing of some eukaryotic tRNA
and rRNA precursors (although these post- transcriptional modifications use a different
splicing mechanism).

A. Eukaryotic mRNA Molecules Have Modified Ends

All eukaryotic mRNA precursors undergo modifications that increase the stability of
the mature mRNAs and make them better substrates for translation. One way to in-
crease the stability of mRNAs is to modify their ends so that they are no longer suscep-
tible to cellular exonucleases that degrade RNA.

The 5' ends are modified while the mRNA precursors are still being synthesized.
The 5' end of the primary transcript is a nucleoside triphosphate residue (usually a
purine) that was the first nucleotide incorporated by RNA polymerase II. Modification
of this end begins when the gamma-phosphate group is removed by the action of a
phosphohydrolase (Figure 21.27). The resulting 5 '-diphosphate group then reacts with
the a-phosphorus atom of a GTP molecule to create a 5' -5' triphosphate linkage. This
reaction is catalyzed by guanylyltransferase and produces a structure called a cap.
The cap is often further modified by methylating the newly added guanylate. The
2 '-hydroxyl groups of the first two nucleotides in the original transcript may also be
methylated. Methyl groups for these reactions are donated by S-adenosylmethionine
(Section 7.3).

The 5' -5' triphosphate linkage protects the mRNA molecule from 5' exonucle-
ases by blocking its 5' end. The cap also converts mRNA precursors into substrates for
other processing enzymes in the nucleus, such as those that catalyze splicing. In mature
mRNA, the cap is the site where ribosomes attach during protein synthesis. Capping is
a cotranscriptional process that is confined to the nucleus. The capping enzymes
shown in Figure 21.27 interact directly with RNA polymerase II transcription com-
plexes but not with RNA polymerase I or RNA polymerase III complexes, ensuring
that mRNA precursors are the only capped RNAs (i.e., tRNA and rRNA are not sub-
strates for capping).

Eukaryotic mRNA precursors are also modified at their 3' ends. Once RNA poly-
merase II has transcribed past the 3' end of the coding region of DNA, the newly
synthesized RNA is cleaved by an endonuclease downstream of a specific site whose
consensus recognition sequence is AAUAAA. This sequence is bound by a cleavage and
polyadenylation specificity factor (CPSF), a protein that also interacts with the endonu-
clease and a polymerase (Figure 21.28). After cleaving the RNA, the endonuclease disso-
ciates and multiple adenylate residues are added to the newly generated 3' end of the
molecule. The addition reactions are catalyzed by poly A polymerase, which adds
adenylate residues using ATP as a substrate. Up to 250 nucleotides can be added to form
a stretch of polyadenylate known as a poly A tail.

With a few rare exceptions, all mature mRNA molecules in eukaryotes contain poly
A tails. The length of the tail varies, depending on the species and possibly on the type
of mRNA and the developmental stage of the cell. The length also depends on the age of
the mRNA since the poly A tail is progressively shortened by the action of 3' exonucle-
ases. In fact, the tail has already been shortened by 50 to 100 nucleotides by the time the
mature mRNA reaches the nuclear pores. The presence of the poly A tail increases the
time required for the exonucleases to reach the coding region.

KEY CONCEPT

Many eukaryotic coding sequences are
interrupted by introns.

658 CHAPTER 21 Transcription and RNA Processing

Figure 21.27 ►

Formation of a cap at the 5' end of a eukary-
otic mRNA precursor. (1) A phosphohydrolase
catalyzes removal of the phosphate group at
the 5' end of the precursor. (2) The 5' end
then receives a GMP group from GTP in a
reaction catalyzed by guanylyltransferase.

(3) The base of the guanylate group is
methylated at N-7. (4) The 2'-hydroxyl
groups of the terminal and the penultimate
ribose groups of the precursor may also be
methylated.

0 G

°0-P=0

°0— P=0 H 2 0 Pi

1 l t

o — ^ s >

°o-p=o

OH OH

°0— P = 0

3'mRNA

21.9 Eukaryotic mRNA Processing 659

(a) Polyadenylation begins when
RNA polymerase II transcription
complex transcribes through a
polyadenylation signal at the 3'
end of an mRNA precursor.

(b) CPSF binds to the consensus
sequence and forms a complex
containing an RNA
endonuclease. The endonuclease
catalyzes cleavage of the
transcript downstream of the
polyadenylation sequence,
forming a new 3' end. Poly A
polymerase can then bind to the
end of the mRNA precursor.

Consensus

sequence

Poly A
polymerase

mRNA

▲ Figure 21.28

Polyadenylation of a eukaryotic mRNA
precursor.

CPSF

ATP

PPi

Poly A
polymerase

B. Some Eukaryotic mRNA Precursors Are Spliced

Splicing is rare in prokaryotes but it is the rule in animals and flowering plants. Internal
sequences that are removed from the primary RNA transcript are called introns.
Sequences that are present in the primary RNA transcript and in the mature RNA mol-
ecule are called exons. The words intron and exon also refer to the regions of the gene
(DNA) that encode corresponding RNA introns and exons. Since DNA introns are tran-
scribed, they are considered part of the gene. The junctions of introns and exons are
known as splice sites since these are the sites where the mRNA precursor is cut and
joined.

Because of the loss of introns, mature mRNA is often a fraction of the size of the
primary transcript. For example, the gene for triose phosphate isomerase from maize
contains nine exons and eight introns and spans over 3400 bp of DNA. The mature

660 CHAPTER 21 Transcription and RNA Processing

(a)

DNA

Triose phosphate
isomerase gene

Site of 3'
cleavage and
polyadenylation

-v

Exon 1 Exon 2 Exon 3 Exon 4

Exon 5

Exon 6 Exon 7 Exon 8 Exon 9

Translated sequence

Transcription

Processing

mRNA 5 m 7 GTP

Mil

A A A A Awv' AAA

Spliced mRNA

(b)

▲ Figure 21.29

Triose phosphate isomerase gene from maize
and the encoded enzyme, (a) Diagram of the
gene showing nine exons and eight introns.
Some exons contain both translated and un-
translated sequences, (b) Three-dimensional
structure of the protein showing the parts of
the protein encoded by each exon.

mRNA, which includes a poly A tail, is only 1050 nucleotides long (Figure 21.29).
The enzyme itself contains 253 amino acid residues.

It used to be thought that there was a correlation between the intron/exon organi-
zation of a gene and the structure of the protein that the gene encodes. According to this
hypothesis, exons encode protein domains and the presence of introns reflects the
primitive organization of the gene. In other words, introns arose early in evolution.
However, as shown in Figure 21.29b, there is no obvious correlation between exons and
protein structure. Most biochemists and molecular biologists now believe that introns
have been inserted at random locations during the evolution of a gene. The “introns
late” hypothesis states that most primitive genes did not have introns and postulates
that introns arose much later during the evolution of eukaryotes.

Introns can vary in length from as few as 42 bp to as many as 10,000 bp (the lower
limit varies with each species; for example, most C. elegans introns are too small to be
accurately spliced in either a vertebrate cell or cell-free extract). The nucleotide se-
quences at splice sites are similar in all mRNA precursors, but the sequence of the rest of
the intron is not conserved. The vertebrate consensus sequences at the two splice sites
are shown in Figure 21.30. Another short consensus sequence is found within the intron
near the 3' end. This sequence, known as the branch site or the branch-point sequence,
also plays an important role in splicing.

21.9 Eukaryotic mRNA Processing 661

Exon

Intron

Exon

* A *

5T/w> G U r AG U y N Y U R A

YYYYYYYYYNCAGG- 3

—

10-40 nucleotides >

5' splice site

Branch site

3' splice site

consensus sequence

The splicing of an mRNA precursor to remove a single intron requires two transes-
terification reactions: one between the 5' splice site and the branch-site adenylate
residue, and one between the 5' exon and the 3' splice site. The products of these two
reactions are (1) the joined exons and (2) the excised intron in the form of a lariat-
shaped molecule. These splicing reactions are catalyzed by a large RNA-protein com-
plex called the spliceosome. The spliceosome helps to not only retain the intermediate
splicing products but also position the splice sites so that the exons can be precisely
joined (Figure 21.31).

The spliceosome is a large, multisubunit complex. It contains over 100 proteins and
five molecules of RNA whose total length is about 5000 nucleotides. These RNA mole-
cules are called small nuclear RNA (snRNA) molecules and are associated with proteins
to form small nuclear ribonucleoproteins, or snRNPs (pronounced “snurps”). snRNPs
are important not only in the splicing of mRNA precursors but also in other cellular
processes.

There are five different types of snRNAs — Ul, U2, U4, U5, and U6. (U stands for
uracil, a common base in these small RNA molecules.) — and a diploid vertebrate nu-
cleus contains more than 100,000 total copies of snRNA. All five snRNA molecules are
extensively base-paired and contain modified nucleotides. Each snRNP contains one or
two snRNAs plus a number of proteins. Some of these proteins are common to all
snRNPs; others are found in only one class of snRNP.

Biochemical experiments in vitro using purified components have led to a sequential
model for spliceosome assembly (Figure 21.32). Spliceosome formation begins when a Ul
snRNP binds to the newly synthesized 5' splice site of the mRNA precursor. This interac-
tion depends on base pairing between the 5' splice site and a complementary sequence
near the 5' end of the Ul snRNA. A U2 snRNP then binds to the branch site of the intron,
forming a stable complex that covers about 40 nucleotides. Next, a U5 snRNP associates
with the y splice site. Finally, a U4/U6 snRNP joins the complex, and all snRNPs are drawn
together to form the spliceosome. Because several groups have now discovered that these
same snRNPs are found preassembled in a much larger complex, prior to splicing, this
pathway may not accurately reflect the splicing cycle in vivo.

Binding of the Ul, U2, and U5 snRNPs to consensus sequences at the 5' splice
site, branch site, and 3' splice site of the intron positions these three interactive sites
properly for the splicing reaction. The spliceosome then prevents the 5' exon from
diffusing away after cleavage and positions it to be joined to the 3' exon. Once a
spliceosome has formed at an intron, it is quite stable and can be purified from cell
extracts.

Since spliceosomes can be observed on nascent transcripts, it is thought that intron
removal is the rate limiting step in RNA processing. Since the spliceosome, which is al-
most as large as a ribosome, is too large to fit through the nuclear pores, the mRNA pre-
cursors are prevented from leaving the nucleus before processing is complete. Once an
intron is excised, the spliceosome gets recycled and will repeat the catalytic cycle on the
next intron it encounters.

Figure 21.31 ►

Intron removal in mRNA precursors. The spliceosome, a multicomponent RNA-protein complex, cat-
alyzes splicing.

[Adapted from Sharp, P. A. (1987). Splicing of messenger RNA precursors. Science 235:766-771.]

◄ Figure 21.30

Consensus sequences at splice sites in verte-
brates. Highly conserved nucleotides are un-
derlined. Y represents a pyrimidine (U or C),
R represents a purine (A or G), and N repre-
sents any nucleotide. The splice sites,
where the RNA precursor is cut and joined,
are indicated by red arrows, and the branch
site is indicated by a black arrow. The intron
is highlighted in blue.

(a) The spliceosome positions the
adenylate residue at the branch
site near the 5' splice site. The
2'-hydroxyl group of the
adenylate attacks the 5' splice site.

(b) The 2'-hydroxyl group is attached
to the 5' end of the intron, and
the newly created 3'-hydroxyl
group of the exon attacks the 3'
splice site.

662 CHAPTER 21 Transcription and RNA Processing

◄ Figure 21.32
Formation of a spliceosome.

(a) As soon as the 5' splice site exits
the transcription complex, a U1
snRNP binds to it.

U1 snRNP

U2 snRNP

(b) Next, a U2 snRNP binds to the
branch site within the intron.

Intron

U4/U6

snRNP

mRNA (exon)

mRNA

(exon)

U5 snRNP

(c) When the 3' splice site emerges
from the transcription complex, a
U5 snRNP binds, and the
complete spliceosome assembles
around a U4/U6 snRNP.

Summary

1. A gene is a sequence of DNA that is transcribed. Housekeeping
genes encode proteins and RNA molecules that are essential for
normal cellular activities.

2. Cells contain several types of RNA, including transfer RNA, ribo-
somal RNA, messenger RNA, and small RNA molecules.

3. DNA-directed RNA synthesis, or transcription, is catalyzed by RNA
polymerase. Ribonucleoside triphosphates are added in nucleotidyl-
group-transfer reactions using a DNA strand as a template.

4. Transcription begins at a promoter sequence and proceeds in the
5' — » 3' direction. A promoter consensus sequence indicates the
nucleotides most commonly found at each position. The cr sub-
unit of E. coli RNA polymerase increases the affinity of the core
polymerase for a promoter and decreases the affinity for nonpro-
moter sequences. During initiation, a transcription bubble forms
and a short stretch of RNA is synthesized. The a subunit dissoci-
ates in the transition from initiation to chain elongation.

Problems 663

5. Transcription termination in E. coli occurs near pause sites, often
when the RNA forms a hairpin structure. Some terminations re-
quire rho , which binds to single- stranded RNA.

6. In eukaryotes, several different RNA polymerases carry out tran-
scription. Transcription factors interact with the promoter and
RNA polymerase to initiate transcription.

7. Some genes are expressed constitutively, but the transcription of
other genes is regulated. Transcription may be regulated by a re-
pressor or an activator. These are often allosteric proteins.

8. Transcription of the three genes of the lac operon is blocked
when lac repressor binds to two operators near the promoter. The
repressor dissociates from the DNA when it binds the inducer

allolactose. Transcription is activated by a complex of cAMP and
CRP (cAMP regulatory protein).

9. RNA transcripts are frequently modified by processing, which in-
cludes the removal, addition, or modification of nucleotide
residues. Primary transcripts of prokaryotic tRNA and rRNA are
processed by nucleolytic cleavage and covalent modification.

10 . Processing of mRNA in eukaryotes includes the addition of a 5'
cap and a 3' poly A tail to protect the molecule from nuclease di-
gestion. In some cases, introns are removed by splicing. The two
transesterification reactions of splicing are catalyzed by the
spliceosome, a complex containing small nuclear ribonucleopro-
teins (snRNPs).

Problems

1 . A bacterial RNA polymerase elongates RNA at a rate of 70 nucleotides
per second, and each transcription complex covers 70 bp of DNA.

(a) What is the maximum number of RNA molecules that can be
produced per minute from a gene of 6000 bp? (Assume that
initiation is not rate limiting.)

(b) What is the maximum number of transcription complexes
that can be bound to this gene at one time?

2. The E. coli genome is approximately 4600 kb in size and contains
about 4000 genes. The mammalian genome is approximately
33 X 10 6 kb in size and contains at most 30,000 genes. An aver-
age gene in E. coli is 1000 bp long.

(a) Calculate the percentage of E. coli DNA that is not transcribed.

(b) Although many mammalian genes are larger than bacterial
genes, most mammalian gene products are the same size as
bacterial gene products. Calculate the percentage of DNA in
exons in the mammalian genome.

3. There are a variety of methods that will allow you to introduce an
intact eukaryotic gene (e.g., the triose phosphate isomerase gene)
into a prokaryotic cell. Would you expect this gene to be properly
transcribed by prokaryotic RNA polymerase? What about the
converse situation, where an intact prokaryotic gene is introduced
into a eukaryotic cell; will it be properly transcribed by a eukary-
otic transcription complex?

4 . Assume that, in a rare instance, a typical eukaryotic triose phos-
phate isomerase gene contains the correct sequences to permit ac-
curate transcription in a prokaryotic cell. Would the resulting
RNA be properly translated to yield the intact enzyme?

5. Describe how the rate of transcription of the lac operon is
affected when E. coli cells are grown in the presence of (a) lactose
plus glucose, (b) glucose alone, and (c) lactose alone.

6. In the promoter of the E. coli lac operon the —10 region has the
sequence 5'-TATGTT-3'. A mutation named UV5 changes this
sequence to 5'-TATAAT-3' (see Figure 21.6). Transcription from
the lac UV5 promoter is no longer dependent on the CRP-cAMP
complex. Why?

7. When /3-3? 2 P4 tATP is incubated with a eukaryotic cell extract
that is capable of transcription and RNA processing, where does
the label appear in mRNA?

8. Unlike DNA polymerase, RNA polymerase does not have proof-
reading activity. Explain why the lack of proofreading activity is
not detrimental to the cell.

9. Mature mRNA from eukaryotic cells is often purified from
other components in the cell with the use of columns contain-
ing oligo (dT) cellulose. These columns contain short segments
of single-stranded deoxyribose thymidylate residues, oligo(dT),
attached to a cellulose matrix. Explain the rationale for use of
these columns to purify mature mRNA from a mixture of
components.

10 . Rifampicin is a semisynthetic compound made from rifamycin
B, an antibiotic isolated from Streptomyces mediterranei.
Rifampicin is an approved anti-mycobacterial drug that is a
standard component of combination regimens for treating tu-
berculosis and staphylococci infections that resist penicillin.
Recent studies have suggested that rifampicin-resistant tuber-
culosis is becoming more common. For example, 2% of sam-
ples from a survey in Botswana were found to be resistant to
the drug. The table below gives some results from wild type
E. coli and E. coli with a single amino acid change in the
(3 subunit of RNA polymerase (Asp to Tyr at amino acid posi-
tion 516) and their growth response to media that contained
rifampicin. [Severinov, K., Soushko, M., Goldfarb, A., and
Nikiforov, V. (1993). Rifampicin region revisited. /. Biol. Chem.
268:14820-14825].

£. coli Rifampicin 0 (fx g/ml)

Wild type <5

Asp51 6Tyr in /3 subunit >50

°Rifampicin concentration at the point of growth arrest of the E. coli.

(a) What is your interpretation of the data?

(b) What role does the [3 subunit have in RNA polymerase?

11. A segment of DNA from the middle of an E. coli gene has the se-
quence below. Write the mRNA sequences that can be produced
by transcribing this segment in either direction.

CCGGCTAAGATCTGACTAGC

12 . Does the definition of a gene given on page 638 5e [first page of
Chapter 2 1 ] apply to the rRNA and tRNA genes whose primary
transcript is shown in Figure 21.26?

664 CHAPTER 21 Transcription and RNA Processing

13. In general, if we know the genomic DNA sequence of a gene we
can reliably predict the nucleotide sequence of the RNA encoded
by that gene. Is this statement also true for tRNAs in prokaryotes?
What about tRNAs in eukaryotes?

14. Assume that a spliceosome assembles at the first intron of the
gene for triose phosphate isomerase in maize (Figure 21.29)
almost as soon as the intron is transcribed (i.e., after about 500
nucleotides of RNA have been synthesized). How long must the
spliceosome be stable if the splicing reaction cannot occur until
transcription terminates? Assume that the rate of transcription by
RNA polymerase II in maize is 30 nucleotides per second.

15. CRP-cAMP represses transcription of the crp gene. Predict the lo-
cation of the CRP-cAMP binding site relative to the promoter of
the crp gene.

16. Why are mutations within an intron of a protein-coding gene
sometimes detrimental?

17. A deletion in one of the introns in the gene for the triose phos-
phate isomerase moves the branch site to a new location seven
nucleotides away from the 3 '-splice acceptor sequence. Will this
deletion have any affect on splicing of the gene?

Selected Readings

General

Alberts, B., Johnson, A., Lewis, J., and Raff, M.
(2007). Molecular Biology of the Cell , 5th ed.
(New York: Garland) .

Krebs, J., Goldstein, L., and Kilpatrick, S. (2009).
Lewin’s Genes X (New York: Jones & Bartlett).

RNA Polymerases and Transcription

Ardehali, M. B., and Lis, J. T. (2009). Tracking rates
of transcription and splicing in vivo. Nature Struc-
tural & Molecular Biology 16:1123-1124.

Bushnell, D. A., and Kornberg, R. D. (2003). Com-
plete, 12-subunit RNA polymerase II and 4.1 -A
resolution: implications for the initiation of tran-
scription. Proc. Natl. Acad. Sci. (U.S.A.)
100:6969-6973.

Kornberg, R. D. (1999). Eukaryotic transcriptional
control. Trends Cell Biol. 9:M46-M49.

Lisser, S., and Margalit, H. (1993). Compilation of
E. coli mRNA promoter sequences. Nucleic Acids
Res. 21:1507-1516.

Murakami, K. S., Masuda, S., Campbell, E. A.,
Muzzin, O., and Darst, S. A. (2002). Structural
basis of transcription initiation: an RNA poly-
merase holoenzyme-DNA complex. Science
296:1285-1290.

Richardson, J. P. (1993). Transcription termina-
tion. Crit. Rev. Biochem. 28:1-30.

Regulation of Transcription

Becker, R B., and Horz, W. (2002). ATP-dependent
nucleosome remodeling. Annu. Rev. Biochem.
71:247-273.

Bushman, F. D. (1992). Activators, deactivators
and deactivated activators. Curr. Biol.

2:673-675.

Fuda, N. J., Behfar, M., and Lis, J. T. (2009). Defin-
ing mechanisms that regulate RNA polymerase II
transcription in vivo. Nature 461:186-192.

Harrison, S. C., and Aggarwal, A. K. (1990). DNA
recognition by proteins with the helix-turn-helix
motif. Annu. Rev. Biochem. 59:933-969.

Jacob, F., and Monod, J. (1961). Genetic
regulatory mechanisms in the synthesis of
proteins. J. Mol. Biol. 3: 318-356.

Kolb, A., Busby, S., Buc, H., Garges, S., and Adhya,
S. (1993). Transcriptional regulation by cAMP and
its receptor protein. Annu. Rev. Biochem.
62:749-795.

Myers, L. C., and Kornberg, R. D. (2000). Mediator
of transcriptional regulation. Annu. Rev. Biochem.
69:729-749.

Pan, Y., Tsai, C.-J., Ma, B., and Nussinov, R. (2009).
How do transcription factors select specific bind-
ing sites in the genome? Nature Structural & Mol-
ecular Biology 16:1118-1120.

Wolfe, A. P., and Guschin, D. (2000). Review: chro-
matin structural features and targets that regulate
transcription./. Struct. Biol. 129:102-122.

Workman, J. L., and Kingston, R. E. (1998). Alter-
ation of nucleosome structure as a mechanism of
transcriptional regulation. Annu. Rev. Biochem.

67: 545-579.

RNA Processing

Apirion, D., and Miczak, A. (1993). RNA process-
ing in prokaryotic cells. BioEssays 15:113-120.

Collins, C. A., and Guthrie, C. (2000). The ques-
tion remains: is the spliceosome a ribozyme?
Nature Struct. Biol. 7: 850-854.

James, B. D., Olsen, G. J., Liu, J., and Pace, N. R.
(1988). The secondary structure of ribonuclease P
RNA, the catalytic element of a ribonucleoprotein
enzyme. Cell 52:19-26.

Jurica, M. S., and Moore, M. J. (2003). Pre-mRNA
splicing: awash in a sea of proteins. Molecular Cell
12:5-14.

McKeown, M. (1993). The role of small nuclear
RNAs in RNA splicing. Curr. Biol. 5:448-454.

Nilsen, T. W. (2003). The spliceosome: the most
complex macromolecular machine in the cell?
BioEssays 25:1 147-1 149.

Proudfoot, N. (2000). Connecting transcription to
messenger RNA processing. Trends Biochem. Sci.
25:290-293.

Shatkin, A. J., and Manley, J. L. (2000). The ends of
the affair: capping and polyadenylation. Nature
Struct. Biol. 7: 838-842.

Wahle, E. (1992). The end of the message: 3 '-end
processing leading to polyadenylated messenger
RNA. BioEssays 14: 1 13-1 18.

o c

_ o

° o o o

° o

o o

° c

o o

Protein Synthesis

W e are now ready to examine the final stage of biological information flow:
the translation of mRNA and the polymerization of amino acids into pro-
teins. The essential features of the biochemistry of protein synthesis were
worked out in the decade between 1955 and 1965. It was clear that there was a genetic
code that had to be used to translate a nucleotide sequence into a sequence of amino
acids. In 1955, Francis Crick proposed that the first step in this process was the attach-
ment of an amino acid to a small adapter RNA. Shortly after that, the adapters, now
known as transfer RNAs, were identified. Ribosomes and the other essential components
of the translation machinery were discovered by fractionating cells and reconstituting
protein synthesis in vitro. Workers in several laboratories demonstrated that messenger
RNA is one of the key intermediates in the flow of information from DNA to protein. By
1961, the most important missing ingredient was the nature of the genetic code.

We begin this chapter with a discussion of the genetic code and tRNA structure.
Next, we examine how mRNA, tRNA, ribosomes, and accessory proteins participate in
protein synthesis. We will also present some examples of the regulation of translation
and post-translational processing.

The results indicate that polyuridylic
acid contains the information for the
synthesis of a protein having many of
the characteristics of poly-L-phenylala-
nine. . . . One or more uridylic acid
residues therefore appear to be the
code for phenylalanine. Whether the
code is of the singlet , triplet , etc v
type has not yet been determined.
Polyuridylic acid seemingly functions
as a synthetic template or messenger
RNA , and this stable , cell-free E. coli
system may well synthesize any pro-
tein corresponding to meaningful in-
formation contained in added RNA.

— M. Nirenberg and H. Matthaei, 1961

22.1 The Genetic Code

George Gamow first proposed the basic structural units of the genetic code. He rea-
soned that since the DNA “alphabet” consists of only four “letters” (A, T, C, and G) and
since these four letters encode 20 amino acids, the genetic code might contain “words,”
or codons, with a uniform length of three letters. Two-letter words constructed from any
combination of the four letters produce a vocabulary of only 16 words (4 2 ), not enough
for all 20 amino acids. In contrast, four-letter words produce a vocabulary of 256 words
(4 4 ), far more than are needed. Three-letter words allow a possible vocabulary of 64
words (4 3 ), more than sufficient to specify each of the 20 amino acids but not excessive.

Top: Escherichia coli ribosome. The ribosome, a complex of RNA and protein, is the site where genetic information is
translated into protein.

665

666 CHAPTER 22 Protein Synthesis

▲ The enigma cryptography machine used by
German armed forces during the Second World
War. This mechanical typewriter permitted
the user to adjust its three large dials to en-
crypt outgoing messages before being sent
by telegraph. The recipients could decode
the message by setting the dials on their
enigma machine to match. This type of en-
cryption is extremely difficult to decipher,
but when Allied forces were able to capture
an intact enigma machine they could listen
in on all their enemy’s transmissions.

The “cracking” of the genetic code began with a chance observation by Marshall
Nirenberg and J. Heinrich Matthaei. They discovered that polyuridylate (poly U) could
direct the synthesis of polyphenylalanine in vitro. By showing that UUU encodes
phenylalanine, they identified the first codon.

Between 1962 and 1965, the rest of the code was deciphered by a number of work-
ers, chiefly Nirenberg and H. Gobind Khorana. Overall, it took ten years of hard work to
learn how mRNA encodes proteins. The development of methods for sequencing genes
and proteins has allowed direct comparison of the primary sequences of proteins with
the nucleotide sequences of their corresponding genes. Each time a new protein and its
gene are characterized, the genetic code is confirmed.

Transfer RNA (tRNA) plays an important role in interpreting the genetic code and
translating a nucleotide sequence into an amino acid sequence. tRNAs are the adapters
between mRNA and proteins. One region of a tRNA molecule is covalently linked to a
specific amino acid, while another region on the same tRNA molecule interacts directly
with an mRNA codon by complementary base pairing. It is this processive joining of the
amino acids specified by an mRNA template that allows the precise synthesis of proteins.

In principle, a genetic code made up of three-letter words can be either overlapping
or nonoverlapping (Figure 22.1). If the codons overlap, then each letter is part of more
than one word and mutating a single letter changes several words simultaneously. For
example, in the sequence shown in Figure 22.1a, each letter is part of three different
words in an overlapping code. One of the advantages of a nonoverlapping code (Figure
22.1b) is that each letter is part of only one word; therefore, mutating a single nu-
cleotide affects only one codon. All living organisms use a nonoverlapping genetic code.

Even with a nonoverlapping code, a sequence can be translated in many different
ways, depending on where translation begins. (We will see later that translation does not
typically begin with the very first nucleotide in an mRNA.) Each potential translation
initiation point defines a unique sequence of three-letter words, or reading frame, in the
mRNA. The correct translation of the “message” transcribed, or written, in the genetic
code depends on establishing the correct reading frame for translation (Figure 22.2).

The standard genetic code is shown in Figure 22.3. With a few minor exceptions, all
living organisms use this genetic code, suggesting that all modern species are descended
from a common ancestor that also used the standard genetic code. This ancestral
species probably lived billions of years ago, making the genetic code one of the most an-
cient remnants of early life.

By convention, all nucleotide sequences are written in the 5' — » 3' direction. Thus,
UAC specifies tyrosine, and CAU specifies histidine. The term codon usually refers to
triplets of nucleotides in mRNA but it can also apply to triplets of nucleotides in the
DNA sequence of a gene. For example, one DNA codon for tyrosine is TAC.

Codons are always translated 5' — > 3', beginning near the 5' end of the message
(i.e., the end synthesized first) and proceeding to the end of the coding region that is

Figure 22.1 ►

Message read in (a) overlapping and
(b) nonoverlapping three-letter codes. In an

overlapping code, each letter is part of three
different three-letter words (as indicated for
the letter G in blue); in a nonoverlapping
code, each letter is part of only one three-
letter word.

mRNA • • • AUGCAUGCAUGC* • •

(a) Message read in
overlapping
triplet code

AUG
U G C
GCA
CAU

(b) Message read in
nonoverlapping
triplet code

AUG

CAU

GCA

U G C

22.1 The Genetic Code 667

usually near the 3' end of the mRNA. The correct reading frame is specified
by special punctuation signals that mark the beginning and the end.

The standard genetic code has several prominent features:

1. The genetic code is unambiguous. In a particular organism or organelle
each codon corresponds to one, and only one, amino acid.

2. There are multiple codons for most amino acids. For example, leucine is
the most abundant amino acid found in proteins (Table 3.3) and has six
codons. Because of the existence of several codons for most amino acids,
the genetic code is said to be degenerate. Different codons that specify the
same amino acid (e.g., UCU and CGU both specify Ser; ACA, ACC, ACG,
and ACU all specify Thr) are known as synonymous codons.

3. The first two nucleotides of a codon are often enough to specify a given
amino acid. For example, the four codons for glycine (GGU, GGC, GGA,
and GGG) all begin with GG.

mRNA -A UGCAUGCAUGC-

Message read in •••a _ UG CaTj gTalTgC--
reading frame 1

Message read in u~^~c a^Tg CAU . .

reading frame 2

Message read in • • -a~U GCA U~GC aTTg C- • ■
reading frame 3

▲ Figure 22.2

One mRNA contains three different reading frames. The

same string of letters read in three different reading
frames will be translated into three different “messages”
or protein sequences. Thus, translation of the correct
message requires selecting the correct reading frame.

4. Codons with similar sequences specify chemically similar amino acids.

For example, the codons for threonine differ from four of the codons for

serine by only a single nucleotide at the 5' position and the codons for aspartate
and glutamate begin with GA and differ only at the 3' position. In addition, codons
with pyrimidines at the second position usually encode hydrophobic amino acids.
Therefore, mutations that alter either the 5' or the 3' position of these codons usu-
ally result in the incorporation of a chemically similar amino acid into the protein.

5. Only 61 of the 64 codons specify amino acids. The three remaining codons (UAA,
UGA, and UAG) are termination codons, or stop codons. Termination codons are not
normally recognized by any tRNA molecules in the cell. Instead, they are recog-
nized by specific proteins that cause newly synthesized peptides to be released from
the translation machinery. The methionine codon, AUG, also specifies the initia-
tion site for protein synthesis and is often called the initiation codon.

Since the completion of the first draft of the human genome in 2000, it has been
common to read in the popular press of “deciphering the code of life” or “unlocking the
human genetic code ” Strictly speaking, the information in the human genome is en-
coded using the same “universal” genetic code discovered 50 years ago. Sequencing projects
actually reveal the messages encoded by the genes and not the code itself.

First position Second position Third position

(5' end)

(3' end)

Phe

Ser

Tyr

Cys

Phe

Ser

Tyr

Cys

Leu

Ser

STOP

Leu

Ser

STOP

Trp

Leu

Pro

His

Arg

Leu

Pro

His

Arg

Leu

Pro

Gin

Arg

Leu

Pro

Gin

Arg

lie

Thr

Asn

Ser

lie

Thr

Asn

Ser

lie

Thr

Lys

Arg

Met

Thr

Lys

Arg

Val

Ala

Asp

Gly

Val

Ala

Asp

Gly

Val

Ala

Glu

Gly

Val

Ala

Glu

Gly

INTERNATIONAL MORSE CODE

Time of dash equals three dots

A • -

N - •

B — • • •

2 . .

D — • •

4 .... —

E •

F .

S • • •

6 — • • • •

T -

7 . . .

H • • • •

u • •-

1 • •

V • • • —

L •

X -• • -
Y

▲ Morse code permitted text to be sent by
telegraph. Messages written in the Latin
alphabet and/or Arabic numerals could be
transmitted via electrical wires using a code
invented by Samuel Morse. In the Morse
code the most common letters in English
language text are coded by the shortest
sequence of dashes and dots (allowing
messages to be sent with the fewest
number of symbols).

◄ Figure 22.3

Standard genetic code. The standard genetic
code is composed of 64 triplet codons. The
left-hand column indicates the nucleotide
found at the first (5') position of the codon;
the top row indicates the nucleotide found
at the second (middle) position of the
codon; and the right column indicates the
nucleotide found at the third (3') position
of the codon. The codon AUG specifies
methionine (Met) and is also used to
initiate protein synthesis. STOP indicates
a termination codon.

668

CHAPTER 22 Protein Synthesis

22.2 Transfer RNA

Transfer RNA molecules are the interpreters of the genetic code. They are the crucial
link between the sequence of nucleotides in mRNA and the sequence of amino acids in
the corresponding polypeptide. In order for tRNA to fulfill this role, every cell must
contain at least 20 different tRNA species (one for every amino acid) and each tRNA
must recognize at least one codon.

A. The Three-Dimensional Structure of tRNA

The nucleotide sequences of different tRNA molecules from many organisms have been
determined. The sequences of almost all these molecules are compatible with the
secondary structure shown in Figure 22.4. This “cloverleaf” structure contains several
arms that are composed of a loop or a loop with a hydrogen-bonded stem. The double-
stranded region of each arm forms a short, stacked, right-handed helix similar to that of
double- stranded DNA.

The 5' end and the region near the 3' end of the tRNA molecule are base-paired to
each other forming the acceptor stem (or amino acid stem). The activated amino acid
will be covalently attached to tRNA on the 3' end of this stem. The amino acid’s car-
boxyl group gets linked to the terminal adenylate’s ribose on either its 2'- or 3 '-hydroxyl
group (Recall from Section 21.8A that mature tRNA molecules are produced by pro-
cessing a larger primary transcript and that the nucleotides at the 3' end of a mature
tRNA molecule are invariably CCA.) All tRNA molecules have a phosphorylated
nucleotide on the 5' end.

The single-stranded loop opposite the acceptor stem in the cloverleaf structure is
called the anticodon loop. It contains the anticodon, the three-base sequence that binds
to a complementary codon in mRNA. The arm of the tRNA molecule that contains the
anticodon is called the anticodon arm. The remaining two arms of the tRNA molecule
are named for the covalently modified nucleotides found within them. (See Figure 21.25

Figure 22.4 ►

Cloverleaf secondary structure of tRNA.

Watson-Crick base pairing is indicated by
dashed lines between nucleotide residues.
The molecule is divided into an acceptor
stem and four arms. The acceptor stem is
the site of amino acid attachment, and the
anticodon arm is the region of the tRNA
molecule that interacts with mRNA codons.
The D and TipC arms are named for modi-
fied nucleotides that are conserved within
these arms. The number of nucleotide
residues in each arm is more or less con-
stant (except in the variable arm). Con-
served bases (gray) and positions of com-
mon modified nucleotides are noted.
Abbreviations other than standard nu-
cleotides: R, a purine nucleotide; Y, a
pyrimidine nucleotide; rrfiA, 1-methyladeny-
late; m 6 A, /V 6 -methyladenylate; Cm,
2'-0-methylcytidylate; D, dihydrouridylate;
Gm, 2'-0-methylguanylate; rrfiG, 1-methyl-
guanylate; m 7 G, 7-methylguanylate; I,
inosinate; i//, pseudouridylate; T, thymine
ribonucleotide.

Acceptor stem
OH

- nrfiA

22.2 Transfer RNA 669

for the structures of these nucleotides.) One of the arms, called the Ti/rC arm, always
contains the triplet sequence ribothymidylate (T), pseudouridylate (i if/), and cytidylate
(C). Dihydrouridylate (D) residues lend their name to the D arm. tRNA molecules also
have a variable arm between the anticodon arm and the Ti/jC arm. The variable arm
ranges in length from about 3 to 21 nucleotides. With a few rare exceptions, tRNA
molecules are between 73 and 95 nucleotides long.

The cloverleaf diagram of tRNA is a two-dimensional representation of a three-
dimensional molecule. In three dimensions, the tRNA molecule is folded into
a sideways £C L” shape (Figures 22.5 and 22.6). The acceptor stem is at one end of the
L-shaped molecule, and the anticodon is located in a loop at the opposite end. The re-
sulting structure is compact and very stable, in part because of hydrogen bonds be-
tween the nucleotides in the D, T^C, and variable arms. This base pairing differs from
normal Watson- Crick base pairing. Most of the nucleotides in tRNA are part of two per-
pendicular stacked helices. The interactions between the adjacent stacked base pairs are
additive and make a major contribution to tRNA stability (analogous to the role of base
stacking interactions in the 3D structure of double-stranded DNA we described in
Section 19.2C).

B. tRNA Anticodons Base-Pair with mRNA Codons

tRNA mediated decoding of the information stored in mRNA molecules requires base-
pairing interactions between tRNA anticodons and complementary mRNA codons. The
anticodon of a tRNA molecule therefore determines where the amino acid attached to
its acceptor stem is added to a growing polypeptide chain. Transfer RNA molecules are
named for the amino acid they carry. For example, the tRNA molecule shown in
Figure 22.6 has the anticodon GAA that binds to the phenylalanine codon UUC. Prior
to protein synthesis, phenylalanine is covalently attached to the acceptor stem of this
tRNA. The molecule is therefore designated tRNA phe .

Much of the base pairing between the codon and the anticodon is governed by the
rules of Watson- Crick base pairing: A pairs with U, G pairs with C, and the strands in the
base-paired region are antiparallel. However, some exceptions to these rules led Francis

Tif/C arm Acceptor stem

▲ Figure 22.5

Tertiary structure of tRNA. The cloverleaf-
shaped molecule shown in Figure 22.4
actually folds up into this three-dimensional
shape. The tertiary structure of tRNA results
from base pairing between the TifjC loop and
the D loop, and two stacking interactions
that (a) align the TipC arm with the acceptor
arm, and (b) align the D arm with the anti-
codon arm. For clarity, only the ribose-
phosphate backbone is shown here.

(a)

Covalent attachment of
activated amino acids
occurs at this site

Base pairing with
mRNA codons involves
these exposed bases

tRNA anticodon
mRNA codon

3' — AAG— 5'

II II III

5' — U U C— 3'
'Phe 7

A 36

a 35

G34

◄ Figure 22.6

Structure of tRNA Phe from the yeast
Saccharomyces cerevisiae. (a) Stick model
showing base pairs and the position of the D
arm (red) relative to the Jif/C arm (green).
Note that there are two double-stranded
RNA helices arrayed at right angles to
each other to form an L-shaped structure.

(b) Diagram showing the complement base-
pairing between tRNA Phe and a phe codon
to generate a double-stranded, antiparallel
RNA helix during decoding.

[NDB TRNA10].

670 CHAPTER 22 Protein Synthesis

m N — H

C-M—

H — N

O |\|

= N

▲ Figure 22.7

Inosinate base pairs. Inosinate (I) is often
found at the 5' (wobble) position of a tRNA
anticodon. Inosinate can form hydrogen
bonds with A, C, or U. This versatility in
hydrogen bonding allows a tRNA carrying
a single anticodon to recognize more than
one synonymous codon.

Table 22.1 Predicted base pairing between the 5' (wobble) position
of the anticodon and the 3 ' position of the codon

Nucleotide at 5' (wobble)
position of anticodon

Nucleotide at 3'
position of codon

A or G

U orC

l a

U, A, or C

°l = Inosinate.

Crick to propose that complementary Watson -Crick base pairing is required for only two
of the three base pairs formed. The codon must form Watson- Crick base pairs with the
3' and middle bases of the anticodon but other types of base pairing are permitted at the
5' position of the anticodon. This alternate pairing suggests that the 5' position is con-
formationally flexible. Crick dubbed this flexibility “wobble” and the 5' position of the
anticodon is sometimes called the wobble position.

Table 22.1 summarizes the allowable base pairs between the wobble position of an
anticodon and the third nucleotide of an mRNA codon. When G is at the wobble posi-
tion, for example, it can pair with either C or U (!). The base at the wobble position of
many anticodons is covalently modified permitting additional flexibility in codon
recognition. For example, in several tRNA molecules, G at the 5' anticodon position is
deaminated at C-2 to form inosinate (I), which can hydrogen-bond with A, C, or U
(Figure 22.7). The presence of I at the 5' position of the anticodon explains why
tRNA Ala with the anticodon IGC can bind to three different codons specifying alanine:
GCU, GCC, and GCA (Figure 22.8).

Wobble allows some tRNA molecules to recognize more than one codon but sev-
eral different tRNA molecules are often required to recognize all synonymous codons.
Different tRNA molecules that can attach to the same amino acid are called isoacceptor
tRNA molecules. The term isoacceptor describes not only tRNA molecules with different
anticodons that are covalently attached to the same activated amino acid but also tRNA
molecules with the same anticodon but different primary sequences. Isoacceptor
tRNAs are identified by Roman numerals or by the codons they recognize (i.e.,
tRNAf 13 , tRNA^ a , or tRNA^ G ).

Genome sequencing data reveal that bacterial genomes encode 30 to 60 different
tRNAs and that eukaryotic genomes have genes for as many as 80 different tRNA mole-
cules. Many of the eukaryotic tRNA genes are present in multiple copies, especially those
genes that encode abundant tRNAs used most frequently in protein synthesis.

22.3 Aminoacyl-tRNA Synthetases

Like DNA and RNA synthesis, protein synthesis can be divided into three distinct
stages: initiation, chain elongation, and termination. However, our description of trans-
lation includes a step prior to the initiation of polymerization, namely, aminoacylation
of tRNA. The activation of amino acids is considered part of the overall translation
process because it is such an important part of the flow of biological information from
nucleic acid to protein.

Each of the 20 amino acids is covalently attached to the 3' end of its respective
tRNA molecules. The product of this reaction is called an aminoacyl-tRNA. The
amino acid is said to be “activated” for subsequent transfer to a growing polypeptide
chain because the aminoacyl-tRNA is a “high-energy” molecule. A specific aminoacyl-
tRNA molecule is identified by naming both the tRNA and the attached amino acid;

22.3 Aminoacyl-tRNA Synthetases

671

for example, aminoacylated tRNA Ala is called alanyl-tRNA Ala . The various enzymes
that catalyze the aminoacylation reaction are called aminoacyl-tRNA synthetases
(e.g., alanyl-tRNA synthetase).

Most species have at least 20 different aminoacyl-tRNA synthetases in each cell since
there are 20 different amino acids. A few species have two different aminoacyl-tRNA syn-
thetases for the same amino acid. Some bacteria don’t have glutaminyl- or asparaginyl-
tRNA synthetases. In these species, the glutaminyl- and asparaginyl-tRNAs are synthe-
sized by modifying glutamate and aspartate residues after they have been covalently
attached to tRNA Gln and tRNA Asn by glutamyl- and aspartyl-tRNA synthetases (Gluta-
mate and aspartate residues that are bound to their proper tRNAs are not modified.)

Although each synthetase is specific for a particular amino acid, it can recognize
many isoacceptor tRNA molecules. For example, there are six codons for serine and sev-
eral different isoacceptor tRNA Ser molecules. All these different tRNA Ser molecules are
recognized by the organism’s single seryl-tRNA synthetase enzyme. The accuracy of
protein synthesis depends on the ability of aminoacyl-tRNA synthetases to catalyze at-
tachment of the correct amino acid to its corresponding tRNA.

A. The Aminoacyl-tRNA Synthetase Reaction

The activation of an amino acid by its specific aminoacyl-tRNA synthetase requires
ATR The overall reaction is:

mRNA 5'

3' 5'

Wobble position

mRNA

3' 5'

Wobble position

Amino Acid + tRNA + ATP > Aminoacyl-tRNA + AMP + PPj (22.1)

The amino acid is covalently attached to the tRNA molecule by the formation of an
ester linkage between the carboxylate group of the amino acid and a hydroxyl group of
the ribose at the 3' end of the tRNA molecule. Since all tRNAs end in — CCA, the at-
tachment site is always an adenylate residue.

Aminoacylation proceeds in two discrete steps (Figure 22.9). In the first step, the
amino acid is activated by formation of a reactive aminoacyl- adenylate intermediate.
The intermediate remains tightly but noncovalently bound to the aminoacyl-tRNA
synthetase. Rapid hydrolysis of the liberated pyrophosphate strongly favors the for-
ward reaction. The second step of aminoacyl-tRNA formation is aminoacyl-group
transfer from the aminoacyl- adenylate intermediate to tRNA. The amino acid is at-
tached to either the 2'- or the 3 '-hydroxyl group of the terminal adenylate residue of
tRNA, depending on the specific aminoacyl-tRNA synthetase catalyzing the reaction.
If the amino acid is initially attached to the 2' -hydroxyl group, it is shifted to the
3 '-hydroxyl group in an additional step. The amino acid must be attached to the 3'
position to function as a protein synthesis substrate.

Formation of the aminoacyl-tRNA is favored under cellular conditions and the intra-
cellular concentration of free tRNA is very low. The Gibbs free energy of hydrolysis of an
aminoacyl-tRNA is approximately equivalent to that of a phosphoanhydride bond in ATP.
The energy stored in the aminoacyl-tRNA is ultimately used in the formation of a peptide
bond during protein synthesis. Note that the two ATP equivalents consumed during each
aminoacylation reaction contribute to the energetic cost of protein synthesis.

▲ Figure 22.8

Base pairing at the wobble position. The

tRNA Ala molecule with the anticodon IGC
can bind to any one of three codons specify-
ing alanine (GCU, GCC, or GCA) because I
can pair with U, C, or A. Note that the RNA
strand containing the codon and the strand
containing the anticodon are antiparallel.
The wobble position is boxed in each
example.

B. Specificity of Aminoacyl-tRNA Synthetases

Attaching a specific amino acid to its corresponding tRNA is a crucial step in translating
a genetic message. If there are errors at this step, the wrong amino acid could be incor-
porated into a protein.

Each aminoacyl-tRNA synthetase binds ATP and selects the proper amino acid
based on its charge, size, and hydrophobicity. This initial selection eliminates most of
the other amino acids. For example, tyrosyl-tRNA synthetase almost always binds tyro-
sine but rarely phenylalanine or any other amino acid. The synthetase then selectively
binds a specific tRNA molecule. The proper tRNA is distinguished by features unique to
its structure. In particular, the part of the acceptor stem that lies on the inner surface of

672 CHAPTER 22 Protein Synthesis

Figure 22.9 ►

Synthesis of an aminoacyl-tRNA molecule
catalyzed by its specific aminoacyl-tRNA
synthetase. In the first step, the nucle-
ophilic carboxylate group of the amino acid
attacks the a-phosphorus atom of ATP,
displacing pyrophosphate and producing an
aminoacyl-adenylate intermediate. In the
second step, nucleophilic attack by the
3'-hydroxyl group of the terminal residue of
tRNA leads to displacement of AMP and for-
mation of an aminoacyl-tRNA molecule.

OH OH

ATP

O 0 O 0 O 0

H 3 N — C — H Amino acid
R

H,0

PPi

Pyrophosphatase

2 P:

STEP 1

OH OH 5'tRNA

( 2 )

AMP

STEP 2

5'tRNA

C=0

H 3 N — C — H
R

3'aminoacyl-tRNA

the L- shaped tRNA molecule is implicated in the binding of tRNA to the aminoacyl-
tRNA synthetase (Figure 22.10).

In some cases, the synthetase enzyme recognizes not only the the acceptor stem of
the tRNA but also the anticodon. For example, the glutaminyl-tRNA synthetase’s ability
to recognize Gln-tRNAs and to discriminate against the other 19 types of tRNAs ensures

22.4 Ribosomes 673

that glutamine is specifically attached to the correct tRNA (shown in Figure
22.10). Note that glutaminyl-tRNA synthetase contacts both the ac-
ceptor stem and the anticodon region of tRNA Gln . The crystal structure
also shows a molecule of ATP bound in the active site near the 3' end of
the tRNA.

Half of the 20 different aminoacyl-tRNA synthetases resemble gluta-
minyl-tRNA synthetase. These enzymes bind the anticodon and aminoacy-
late tRNA at the 2 '-hydroxyl group. A subsequent chemical rearrangement
shifts the aminoacyl group to the 3 '-hydroxyl group. Such enzymes are
known as class I synthetases. Class II aminoacyl-tRNA synthetases are
often more complex, multisubunit enzymes and they aminoacylate tRNA
at the 3 '-hydroxyl group. In all cases, the net effect of the interaction be-
tween tRNA and synthetase is to position the 3' end of the tRNA molecule
in the active site of the enzyme.

C. Proofreading Activity of Aminoacyl-tRNA Synthetases

The error rate for most aminoacyl-tRNA synthetases is low because they
make multiple contacts with a specific tRNA and a specific amino acid.

However, isoleucine and valine are chemically similar amino acids, and
both can be accommodated in the active site of isoleucyl-tRNA synthetase
(Figure 22.11). Isoleucyl-tRNA synthetase mistakenly catalyzes the forma-
tion of the valyl- adenylate intermediate about 1% of the time. On the basis
of this observation, we might expect valine to be attached to isoleucyl-tRNA and incorpo-
rated into protein in place of isoleucine about 1 time in 100 but the observed substitution
of valine for isoleucine in polypeptide chains is only about 1 time in 10,000. This lower
level of valine incorporation suggests that isoleucyl-tRNA synthetase also discriminates
between the two amino acids after aminoacyl- adenylate formation. In fact, isoleucyl-
tRNA synthetase carries out proofreading in the next step of the reaction. Although
isoleucyl-tRNA synthetase may mistakenly catalyze the formation of valyl- adenylate, it
usually catalyzes hydrolysis of the incorrect valyl-adenylate to valine and AMP or the hy-
drolysis of valyl-tRNA 1 . The overall error rate of the reaction is 10 -5 for most amino
acyl-tRNA synthetases.

22.4 Ribosomes

Acceptor stem

Anticodon

▲ Figure 22.10

Structure of E. co//tRNA Gln bound to gluta-
minyl-tRNA synthetase. The 3' end of the

tRNA is buried in a pocket on the surface of
the enzyme. A molecule of ATP is also
bound at this site. The enzyme interacts
with both the tRNA acceptor stem and anti-
codon. [PDB 1QRS].

KEY CONCEPT

The accuracy of information flow from nu-
cleic acids to protein depends, in part, on
the accuracy of the amino acyl-tRNA
synthetase reaction.

Protein synthesis requires assembling four components that form an elaborate translation
complex: the ribosome, which catalyzes peptide bond formation; its accessory protein fac-
tors, which help the ribosome in each step of the process; the mRNA, which carries the in-
formation specifying the protein s sequence; and the aminoacyl-tRNAs that carry the acti-
vated amino acids. Initiation involves assembly of the translation complex at the first
codon in the mRNA. During polypeptide chain elongation the ribosome and associated
components move, or translocate, along the template mRNA in the 5' — » 3' direction.

◄ Figure 22.11

Model of the substrate-binding site in
isoleucyl-tRNA synthetase. Despite the
similar size and charge of isoleucine and
valine, isoleucyl-tRNA synthetase binds to
isoleucine about 100 times more readily
than it binds to valine. A subsequent proof-
reading step also helps prevent the forma-
tion of valyl-tRNA lle .

674

CHAPTER 22 Protein Synthesis

The polypeptide is synthesized from the N- terminus to its C- terminus. Finally, when syn-
thesis of the protein is complete, the translation complex disassembles in a separate termi-
nation step. An important function of disassembly is to release the two ribosomal sub-
units from the mRNA so that they can participate in further rounds of translation.

A. Ribosomes Are Composed of Both Ribosomal RNA and Protein

All ribosomes contain two subunits of unequal size. In E. coli, the small subunit is called
the 30S subunit and the large subunit is called the 50S subunit. (The terms 30S and 50S
originally referred to the sedimentation rate of these subunits.) The 30S subunit is elon-
gated and asymmetric, with overall dimensions of 5.5 X 22 X 22.5 nm. A narrow neck
separates the head from the base and a protrusion extends from the base forming a cleft
where the mRNA molecule appears to rest. The 50S ribosomal subunit is wider than the
30S subunit and has several protrusions; its dimensions are about 15 X 20 X 20 nm.
The 50S subunit also contains a tunnel about 10 nm long and 2.5 nm in diameter. This
tunnel extends from the site of peptide bond formation and accommodates the growing
polypeptide chain during protein synthesis. The 30S and 50S subunits combine to form
an active 70S ribosome.

In E. coli, the RNA component of the 30S subunit is a 16S rRNA of 1542 nu-
cleotides. Although its exact length varies among species, the 16S rRNA contains exten-
sive regions of secondary structure that are highly conserved in the ribosomes of all
living organisms. There are 21 ribosomal proteins in the 30S subunit. The 50S subunit
of the E. coli ribosome contains two molecules of ribosomal RNA: one 5S rRNA of 120
nucleotides and one 23S rRNA of 2904 nucleotides. There are 31 different proteins asso-
ciated with the 5S and 23S rRNA molecules in the 50S subunit (Figure 22.12).

Eukaryotic ribosomes are similar in shape to bacterial ribosomes but they tend to be
somewhat larger and more complex. Intact vertebrate ribosomes are designated 80S and
are made up of 40S and 60S subunits (Figure 22.12). The small 40S subunit is analogous
to the 30S subunit of the prokaryotic ribosome; it contains about 30 proteins and a single
molecule of 18S rRNA. The large 60S subunit contains about 40 proteins and three riboso-
mal RNA molecules: 5S rRNA, 28S rRNA, and 5.8S rRNA. The 5.8S rRNA is about 160
nucleotides long and its sequence is similar to that of the 5' end of prokaryotic 23S
rRNA. This similarity implies that the 5.8S rRNA and the 5' end of prokaryotic 23S

5 '

50S

23S

rRNA

3 '

5S rRNA
31 proteins

70S

-> 21 proteins
■» 16S rRNA

30 proteins
18S rRNA

Prokaryote

Eukaryote

▲ Figure 22.12

Comparison of prokaryotic and eukaryotic ribosomes. Both types of ribosomes consist of two subunits, each of which contains ribosomal RNA and
proteins. The large subunit of the prokaryotic ribosome contains two molecules of rRNA: 5S and 23S. The large subunit of almost all eukaryotic ribo-
somes contains three molecules of rRNA: 5S, 5.8S, and 28S. The sequence of the eukaryotic 5.8S rRNA is similar to the sequence of the 5' end of
the prokaryotic 23S rRNA.

22.5 Initiation of Translation 675

rRNA are derived from a common ancestor and that there has been a fusion or splitting
of rRNA genes during their evolution.

Both prokaryotic and eukaryotic genomes contain multiple copies of ribosomal RNA
genes. The combination of a large number of copies and strong promoters for these genes
allows cells to maintain a high level of ribosome synthesis. Eukaryotic ribosomal RNA
genes, which are transcribed by RNA polymerase I (Section 21.5A), occur as tandem arrays
of hundreds of copies. In most eukaryotes, these genes are clustered in the nucleolus, where
processing of ribosomal RNA precursors and ribosome assembly occur (Section 21.8B).
This processing is coupled to ribosome assembly, as shown in Figure 22.13 for the E. coli
30S subunit. Many of the ribosomal proteins contact RNA and bind specifically to regions
of secondary structure in 16S rRNA. Others form protein-protein contacts and assemble
into the complex only when other ribosomal proteins are present.

The structure of the 30S ribosomal subunit from the bacterium Thermus ther-
mophilus is shown in Figure 22.14 on page 676. Note that most of the mass of the 30S
subunit is due to the 16S ribosomal RNA, which forms a compact structure made up of
multiple regions of double-stranded RNA. The ribosomal proteins bind to the surface of
the RNA or to grooves and crevices between regions of RNA secondary structure.

Similarly, the assembly of the bacterial 50S subunit and of the 40S and 60S eukary-
otic subunits are also coupled to the processing of their ribosomal RNA precursors. The
structure of the 50S subunit from the archeon Haloarcula marismortui is also shown in
Figure 22.14.

B. Ribosomes Contain Two Aminoacyl-tRNA Binding Sites

As discussed in Section 22.3, the substrates for peptide bond formation are not free
amino acids but relatively large aminoacyl-tRNA molecules. A ribosome must align two
adjacent aminoacyl-tRNA molecules so that their anticodons interact with the correct
mRNA codons. The aminoacylated ends of these two tRNAs are positioned at the site of
peptide bond formation. The ribosome must also hold the mRNA and the growing
polypeptide chain, and it must accommodate the binding of several protein factors dur-
ing protein synthesis. The ability to accomplish these tasks simultaneously explains, in
part, why the ribosome is so large and complex.

The orientation of the two tRNA molecules during protein synthesis is shown in
Figure 22.15 on page 677. The growing polypeptide chain is covalently attached to the
tRNA positioned at the peptidyl site (P site), forming peptidyl-tRNA. The second
aminoacyl-tRNA is bound at the aminoacyl site (A site). As the polypeptide chain is
synthesized, it passes through the tunnel of the large ribosomal subunit and emerges on
the outer surface of the ribosome.

22.5 Initiation of Translation

The initiation of protein synthesis involves assembling a translation complex at the begin-
ning of an mRNAs coding sequence. This complex consists of the two ribosomal subunits,
an mRNA template to be translated, an initiator tRNA molecule, and several accessory pro-
teins called initiation factors. This crucial initiation step ensures that the proper initiation
codon (and therefore the correct reading frame) is selected before translation begins.

A. Initiator tRNA

As mentioned in Section 22.1, the first codon translated is usually AUG. Every cell con-
tains at least two types of methionyl-tRNA Met molecules that can recognize AUG
codons. One type is used exclusively at initiation codons and is called the initiator
tRNA. The other type only recognizes internal methionine codons. Although these two
tRNA Met molecules have different primary sequences, and distinct functions, both of
them are aminoacylated by the same methionyl-tRNA synthetase.

In bacteria, the initiator tRNA is called tRNAf 161 . The charged initiator tRNA
(methionyl-tRNAf 161 ) is the substrate for a formyltransferase that catalyzes addition of
a formyl group from 10-formyltetrahydrofolate to the methionine residue producing

21S particle

Complete 30S subunit
▲ Figure 22.13

Assembly of the 30S ribosomal subunit and
maturation of 16S rRNA in E. coli. Assembly
of the 30S ribosomal subunit begins when
six or seven ribosomal proteins bind to the
16S rRNA precursor as it is being tran-
scribed, thereby forming a 21S particle.

The 21S particle undergoes a conforma-
tional change, and the 16S rRNA molecule
is processed to its final length. During this
processing, the remaining ribosomal pro-
teins of the 30S subunit bind (recall that
M16 is a site-specific endonuclease in-
volved in RNA processing that we discussed
in Chapter 21).

676 CHAPTER 22 Protein Synthesis

Central Protuberance

50S subunit interface
(Crown View)

L7/L12 Stalk

Domain IV
Ridge

Shoulder

Protein
S12

Body

Head

Neck

Platform

Spur

Helix 44

Central Protuberance

180°

50S solvent face

Head

Neck

Shoulder

Platform

Spur

30S subunit interface

30S solvent face

▲ Figure 22.14

Three-dimensional structures of the
H. marismortui 50S subunit (top) and the
T. thermophilus 30S subunit (bottom).

N-formylmethionyl-tRNA^^fMet-tRNAf 1 ^) as shown in Figure 22.16 on page 681. In
eukaryotes and archaebacteria, the initiator tRNA is called tRNA- 461 . The methionine
that begins protein synthesis in eukaryotes is not formylated.

N-Formylmethionine in bacteria — or methionine in other organisms — is the
first amino acid incorporated into proteins. After protein synthesis is under way, the
N- terminal methionine can be either deformylated or removed from the polypeptide
chain altogether.

B. Initiation Complexes Assemble Only at Initiation Codons

There are three possible reading frames in an mRNA molecule but only one of them is
correct. Establishing the correct reading frame during the initiation of translation is

22.5 Initiation of Translation

677

critical for the accurate decoding of information from mRNA into protein. Shifting
the reading frame by even a single nucleotide would alter the sequence of the entire
polypeptide and result in a nonfunctional protein. The translation machinery must
therefore accurately locate the initiation codon that serves as the start site for protein
synthesis.

The ribosome needs to distinguish between the single correct initiation codon
and all the other incorrect AUGs. These other AUGs specify either internal methion-
ine residues in the correct reading frame or irrelevant methionine codons in the two
other incorrect reading frames. It is important to appreciate that the initiation codon
is not simply the first three nucleotides of the mRNA. Initiation codons can be lo-
cated many nucleotides downstream of the 5 '-end of the mRNA molecule.

In prokaryotes, the selection of an initiation site depends on an interaction be-
tween the small subunit of the ribosome and the mRNA template. The 30S subunit
binds to the mRNA template at a purine- rich region just upstream of the correct initia-
tion codon. This region, called the Shine- Dalgarno sequence, is complementary to a
pyrimidine- rich stretch at the 3' end of the 16S rRNA molecule. During formation of the
initiation complex, these complementary nucleotides pair to form a double-stranded
RNA structure that binds the mRNA to the ribosome. The result of this interaction is to
position the initiation codon at the P site on the ribosome (Figure 22.17). The initiation
complex assembles exclusively at initiation codons because Shine-Dalgarno sequences are
not found immediately upstream of internal methionine codons.

70S ribosome

P site

A site

tRNA with
amino acid

mRNA
5'

Tunnel

Growing

peptide

chain

▲ Figure 22.15

Sites for tRNA binding in prokaryotic ribo-
somes. During protein synthesis, the P site
is occupied by the tRNA molecule attached
to the growing polypeptide chain, and the A
site holds an aminoacyl-tRNA. The growing
polypeptide chain passes through the tunnel
of the large subunit.

C. Initiation Factors Help Form the Initiation Complex

Formation of the initiation complex requires several initiation factors in addition to
ribosomes, initiator tRNA, and mRNA. Prokaryotes contain three initiation factors,
designated IF- 1, IF-2, and IF-3. There are at least eight eukaryotic initiation factors
(elF’s). In both prokaryotes and eukaryotes, the initiation factors catalyze assembly of
the protein synthesis complex at the initiation codon.

(a)

Lipoprotein

— AU CUAGAGGGU

RecA

— G G C AUG A C AGG

GalE

—AG CCUAAUGGA

GalT

— C C CGAUUAAGG

Lacl

— CAAUUCAGGG U

LacZ

— U U CACAC AGGA

Ribosomal L10

— CAUCAAGGAGC

Ribosomal L7/L12

— UAUUCAGGAAC

I 1 I I I I I I

AUUAAUAAUGAAAGCUACU—

AGUAAAAAUGGCUAUCG—

GCGAAUUAUGAGAGUUCUG—

AACGACCAUGACGCAAUUU—

GGUGAAUGUGAAACCAGUA—

AACAGCUAUGACCAUGAUU—

AAAGCUAAUGGCUUUAAAU—

AAUUUAAAUGU CUAUCACU—

tRNA f Met

c=o

c— N— c— H
I I

H CH 2

ch 2

ch 3

▲ Figure 22.16

Chemical structure of fMet-tRNA f Met . A

formyl group (red) is added to the
methionyl moiety (blue) of methionyl-
tRNA f Met in a reaction catalyzed by a
formyltransferase.

(b)

3'end of 16S rRNA

hHO C U'

X A A^"

u uccuc c

fMet Thr Met lie

dJ UCACAC AGGAAACAGCU AUGACCAUGAU U— mRNA

, , , 3 '

Shine-Dalgarno ! ! !

sequence ACv^

Anticodon
of fMet-tRNA f Met

◄ Figure 22.17

Shine-Dalgarno sequences in E. coli mRNA.

(a) Ribosome-binding sites at the 5' end
of mRNA for several E. coli proteins. The
Shine-Dalgarno sequences (red) occur im-
mediately upstream of initiation codons
(blue), (b) Complementary base pairing
between the 3' end of 16S rRNA and the
region near the 5' end of an mRNA. Binding
of the 3' end of the 16S rRNA to the Shine-
Dalgarno sequence helps establish the
correct reading frame for translation by
positioning the initiation codon at the
ribosome’s P site.

678

CHAPTER 22 Protein Synthesis

One of the roles of IF- 3 is to maintain the ribosomal subunits in their dissociated
state by binding to the small subunit. The ribosomal subunits bind separately to the ini-
tiation complex and the association of IF- 3 with the 30S subunit prevents the 30S and
50S subunits from forming the 70S complex prematurely. IF-3 also helps position
fMet-tRNAf 161 and the initiation codon at the P site of the ribosome. IF-2 selects the
initiator tRNA from the pool of aminoacylated tRNA molecules in the cell. It binds
GTP forming an IF-2-GTP complex that specifically recognizes the initiator tRNA and
rejects all other aminoacyl-tRNA molecules. The third initiation factor, IF- 1, binds to
the 30S subunit and facilitates the actions of IF-2 and IF-3.

Once the 30S complex has been formed at the initiation codon, the 50S ribosomal
subunit binds to the 30S subunit. Next, the GTP bound to IF-2 is hydrolyzed and Pj is
released. The initiation factors dissociate from the complex when GTP is hydrolyzed.
IF-2-GTP is regenerated when the bound GDP is exchanged for GTP. The steps in the for-
mation of the 70S initiation complex are summarized in Figure 22.18.

(1) IF-3 and IF-1 bind to the
30S subunit, preventing
premature assembly of the
70S complex.

IF-1

IF-3

v Figure 22.18

Formation of the prokaryotic 70S initiation
complex.

30S subunit

70S initiation
complex

GDP

F-2

IF-3

30S initiation complex

50S subunit

(3) The 50S subunit then joins
the 30S initiation complex,
IF-1 and IF-3 are released,
and the GTP bound to IF-2 is
hydrolyzed to GDP and Pj.
IF-2-GDP dissociates,
leaving the 70S initiation^ 161
complex with fMet-tRNA
positioned in the P site.

GTP

(2) IF-2-GTP binds to the 30S
subunit and facilitates
binding of Met-tRNA^

The 30S complex interacts
with mRNA by recognizing
the Shine-Dalgarno sequence
and the initiation codon.

IF-2

mRNA

fMet-tRNAf

22.6 Chain Elongation During Protein Synthesis Is a Three-Step Microcycle

679

The role of the prokaryotic initiation factors is to ensure that the aminoacylated
initiator tRNA (fMet-tRNAf 401 ) is correctly positioned at the initiation codon. The
initiation factors also mediate the formation of a complete initiation complex by re-
constituting a 70S ribosome such that the initiation codon is positioned in the P site.

D. Translation Initiation in Eukaryotes

Eukaryotic mRNAs do not have distinct Shine-Dalgarno sequences that serve as ribo-
some binding sites. Instead, the first AUG codon in the message usually serves as the ini-
tiation codon. eIF-4 (eukaryotic initiation factor 4), also known as cap binding protein
(CBP), binds specifically to the 7-methylguanylate cap (Figure 21.26) at the 5' end of
eukaryotic mRNA. Binding of eIF-4 to the cap structure leads to the formation of a
preinitiation complex consisting of the 40S ribosomal subunit, an aminoacylated initia-
tor tRNA, and several other initiation factors. The preinitiation complex then scans
along the mRNA in the 5' — > 3' direction until it encounters an initiation codon. When
the search is successful, the small ribosomal subunit is positioned so that Met-tRNAi Met
interacts with the initiation codon in the P site. In the final step, the 60S ribosomal sub-
unit binds to complete the 80S initiation complex and all the initiation factors dissoci-
ate. The dissociation of eIF-2 — the eukaryotic counterpart of bacterial IF-2 — is accom-
panied by GTP hydrolysis.

Most eukaryotic mRNA molecules encode only a single polypeptide since the nor-
mal mechanism of selecting the initiation codon by scanning along the mRNA from the
5' end permits only one initiation codon per mRNA. In contrast, prokaryotic mRNAs
often contain several coding regions. Each coding region begins with an initiation
codon that is associated with its own upstream Shine-Dalgarno sequence. mRNA mole-
cules that encode several polypeptides are said to be polycistronic.

22.6 Chain Elongation During Protein Synthesis
Is a Three-Step Microcycle

At the end of the initiation step, the mRNA is positioned so that the next codon can be
translated during the elongation stage of protein synthesis. The initiator tRNA occupies
the P site in the ribosome and the A site is ready to receive an incoming aminoacyl-
tRNA. During chain elongation each additional amino acid is added to the nascent
polypeptide chain in a three-step microcycle. The steps in this microcycle are (1) posi-
tioning the correct aminoacyl-tRNA in the A site of the ribosome, (2) forming the peptide
bond, and (3) shifting, or translocating, the mRNA by one codon relative to the ribo-
some (the two tRNAs in the ribosome’s P and A sites also translocate).

The translation machinery works relatively slowly compared to the enzyme systems
that catalyze DNA replication. Proteins are synthesized at a rate of only 18 amino acid
residues per second, whereas bacterial replisomes synthesize DNA at a rate of 1000
nucleotides per second. This difference in rates reflects, in part, the difference between
polymerizing four types of nucleotides to make nucleic acids and polymerizing 20 types
of amino acids to make proteins. Testing and rejecting all of the incorrect aminoacyl-
tRNA molecules also takes time and slows protein synthesis.

The rate of transcription in prokaryotes is approximately 55 nucleotides per second.
This corresponds to about 18 codons per second or the same rate at which the mRNA
is translated. In bacteria, translation initiation occurs as soon as the 5' end of an
mRNA is synthesized and translation and transcription are coupled (Figure 22.19 on
page 680). This tight coupling is not possible in eukaryotes because transcription and
translation are carried out in separate compartments of the cell (the nucleus and the
cytoplasm, respectively). Eukaryotic mRNA precursors must be processed in the
nucleus (e.g., capped, polyadenylated, spliced) before they are exported to the cyto-
plasm for translation.

An E. coli cell contains about 20,000 ribosomes. Many large eukaryotic cells have
several hundred thousand ribosomes. Farge mRNA molecules can be translated simul-
taneously by many protein synthesis complexes forming a polyribosome or polysome, as

KEY CONCEPT

The A site of an actively translating
ribosome spends the vast majority of its
time bound to one of the 19 types of
incorrect aminoacyl-tRNAs as it randomly
samples the pool of charged tRNAs,
seeking the correct tRNA.

680

CHAPTER 22 Protein Synthesis

Strand of DNA
being transcribed

A polyribosome,
or polysome

Individual ribosomes
synthesizing new proteins
from the mRNAs

▲ Figure 22.19

Coupled transcription and translation of an
E. coli gene. The gene is being transcribed
from left to right. Ribosomes bind to the
5' end of the mRNA molecules as soon as
they are synthesized. The large polysomes
on the right are released from the gene
when transcription terminates.

seen in Figure 22.19. The number of ribosomes bound to an mRNA molecule depends on
the length of the mRNA and the efficiency of initiation of protein synthesis. At maximal
efficiency the spacing between each translation complex in the polysome is about 100
nucleotides. On average, each mRNA molecule in an E. coli cell is translated 30 times,
effectively amplifying the information it encodes by 30-fold.

EF-Tu

tRNA phe

▲ Figure 22.20

EF-Tu binds aminoacylated tRNAs. The EF-

Tu-GTP complex binds to the acceptor
end of aminoacylated tRNA (in this case
phenylalanyl-tRNA Phe ). The phenylalanine
residue is shown in green. This is how
charged tRNAs commonly exist inside a cell.

A. Elongation Factors Dock an Aminoacyl-tRNA in the A Site

At the start of the first chain elongation microcycle, the A site is empty and the P site is
occupied by the aminoacylated initiator tRNA. The first step in chain elongation is in-
sertion of the correct aminoacyl-tRNA into the A site of the ribosome. In bacteria, this
step is catalyzed by an elongation factor called EF-Tu. EF-Tu is a monomeric
protein that contains a binding site for GTR Each E. coli cell has about 135,000
molecules of EF-Tu, making it one of the most abundant proteins in the cell
(emphasizing the importance of protein synthesis to a cell).

EF-Tu-GTP associates with an aminoacyl-tRNA molecule to form a ter-
nary complex that fits into the A site of a ribosome. Almost all aminoacyl-
tRNA molecules in vivo are found in such ternary complexes (Figure 22.20).
The structure of EF-Tu is similar to that of IF-2 (which also binds GTP) and
other G proteins (Section 9.12A), suggesting that they all evolved from a com-
mon ancestral protein.

The EF-Tu-GTP complex recognizes common features of the tertiary
structure of tRNA molecules and binds tightly to all aminoacyl-tRNA mole-
cules except fMet-tRNAf 161 . The fMet-tRNAf 101 molecule is distinguished
from all other aminoacyl-tRNA molecules by the distinctive secondary struc-
ture of its acceptor stem.

A ternary complex of EF-Tu-GTP-aminoacyl-tRNA can diffuse freely into
the A site in the ribosome. When correct base pairs form between the anti-
codon of the aminoacyl-tRNA and the mRNA codon in the A site, the complex
is stabilized. EF-Tu-GTP can then contact sites in the ribosome as well as the
tRNA in the P site (Figure 22.21, on page 681). These contacts trigger hydroly-
sis of GTP to GDP and Pj causing a conformational change in EF-Tu-GDP that
releases the bound aminoacyl-tRNA. EF-Tu-GDP then dissociates from the
chain elongation complex. The aminoacyl-tRNA remains in the A site where it
is positioned for peptide bond formation.

EF-Tu-GDP cannot bind another aminoacyl-tRNA molecule until GDP
dissociates. An additional elongation factor called EF-Ts catalyzes the exchange
of bound GDP for GTP (Figure 22.22, on page 682). Note that one GTP mol-
ecule is hydrolyzed for every aminoacyl-tRNA that is successfully inserted into
the A site.

22.6 Chain Elongation During Protein Synthesis Is a Three-Step Microcycle 681

GDP

A site
occupied

Formation of the correct
complex triggers hydrolysis
of GTP, which alters the
conformation of EF-Tu.
EF-Tu dissociates, leaving
behind a correctly inserted
aminoacyl-tRNA.

◄ Figure 22.21

Insertion of an aminoacyl-tRNA by EF-Tu during
chain elongation in E. coli.

Peptidyl-tRNA
occupies P site

Ternary complex

Anticodon pairs
with codon

Aminoacyl-tRNA

GTP

The ternary complex enters
the A site. If the codon and
anticodon match, EF-Tu forms
contacts with the ribosome and
the peptidyl-tRNA in the P site.

A site

unoccupied

B. Peptidyl Transferase Catalyzes Peptide Bond Formation

Binding of a correct aminoacyl-tRNA in the A site aligns the activated amino acid’s
a -amino group next to the ester bond’s carbonyl on the peptidyl-tRNA in the neighboring
P site. The nitrogen atom’s lone pair of electrons execute a nucleophilic attack on the car-
bonyl carbon, resulting in the formation of a peptide bond via a displacement reaction.
While it is straightforward to visualize how the ribosome’s active site aligns these substrates,
we do not understand precisely how the ribosome enhances the rate of this reaction. The
peptide chain, now one amino acid longer, is transferred from the tRNA in the P site to the
tRNA in the A site (Figure 22.23, on page 683). Formation of the peptide bond requires hy-
drolysis of the energy- rich peptidyl-tRNA linkage. Note that the growing polypeptide chain
is covalently attached to the tRNA in the A site, forming a peptidyl-tRNA.

The enzymatic activity responsible for formation of the peptide bond is referred to
as peptidyl transferase. This activity is contained within the large ribosomal subunit. Both
the 23S rRNA molecule and the 50S ribosomal proteins contribute to the substrate bind-
ing sites, but the catalytic activity is localized to the RNA component. Thus, peptidyl
transferase is yet another example of an RNA- catalyzed reaction.

KEY CONCEPT

Formation of the new peptide bond involves
physically transferring the polypeptide
attached to the P site tRNA onto the amino-
terminus of the aminoacyl-tRNA bound in
the ribosome’s A site.

682 CHAPTER 22 Protein Synthesis

Figure 22.22 ►
Cycling of EF-Tu-GTP.

EF-Tu-GTP-aminoacyl-tRNA

complex

(1) Aminoacyl-tRNA is
delivered to the
ribosome, and GTP is
hydrolyzed, causing the
EF-Tu-GDP complex
to dissociate.

Aminoacyl-tRNA

(4) Regenerated
EF-Tu-GTP
binds another
aminoacyl-
tRNA molecule

EF-Tu-GTP

complex

(3) The EF-Tu-EF-Ts
complex binds GTP,
which causes EF-Ts
to dissociate.

GTP

EF-Tu-GDP

complex

(2) The inactive EF-Tu-GDP
complex is recognized by
elongation factor EF-Ts,
which promotes
dissociation of GDP.

GDP EF-Tu-EF-Ts
complex

C. Translocation Moves the Ribosome by One Codon

After the peptide bond has formed, the newly created peptidyl-tRNA is partially in the
A site and partially in the P site (Figure 22.24, on page 684). The deaminoacylated tRNA
has been displaced somewhat from the P site. It now occupies a position on the ribo-
some that is referred to as the exit site, or E site. Before the next codon can be translated,
the deaminoacylated tRNA must be released and the peptidyl-tRNA must be completely
transferred from the A site to the P site. At the same time, the mRNA must shift by one
codon relative to the ribosome. This translocation is the third step in the chain elonga-
tion microcycle.

In prokaryotes, the translocation step requires a third elongation factor, EF-G. Like
the other elongation factors, EF-G is an abundant protein; an E. coli cell contains
approximately 20,000 molecules of EF-G, or roughly one for every ribosome. Like EF-
Tu, EF-G has a binding site for GTP. Binding of EF-G-GTP to the ribosome completes
the translocation of the peptidyl-tRNA from the A site to the P site and releases the
deaminoacylated tRNA from the E site. EF-G itself is released from the ribosome only
when its bound GTP is hydrolyzed to GDP and Pi is released. The dissociation of EF-
G-GDP leaves the ribosome free to begin another microcycle of chain elongation.

The growing polypeptide chain extends from the peptidyl-tRNA in the
P site through a tunnel in the 50S subunit, to exit on the exterior surface of the ribosome

22.6 Chain Elongation During Protein Synthesis Is a Three-Step Microcycle 683

P site A site

tRNA tRNA

O O

NH R n+2

o=c

H-C-R n

Peptidyl transferase

◄ Figure 22.23

Formation of a peptide bond. The carbonyl
carbon of the peptidyl-tR N A undergoes nu-
cleophilic attack by the nitrogen atom of the
amino group. This aminoacyl-group-transfer
reaction results in growth of the peptide
chain by one residue and transfer of the
nascent peptide to the tRNA in the A site.

P site

A site

tRNA

H — C —

Rn+2

c = o

H-C-R n+1

0 = C

H-C-

684

CHAPTER 22 Protein Synthesis

Peptidyl

transferase

Unoccupied

▲ Figure 22.24

Translocation during protein synthesis in
prokaryotes.

top: Aminoacyl-tRNA is positioned in the
A site.

middle: Following synthesis of the peptide
bond, the newly formed peptidyl-tR N A is
partly in the A site and partly in the P site.

bottom: Translocation shifts the peptidyl-
tRNA completely into the P site, leaving the
A site empty and ejecting the deamino-
acylated tRNA from the E site.

(Figure 22.15). Each translocation step helps push the chain through the tunnel. The
newly synthesized polypeptide doesn’t begin to fold into its final shape until it emerges
from the tunnel. This folding is assisted by chaperones, such as HSP70, that are associ-
ated with the translation machinery (Section 4.10D).

The elongation microcycle is repeated for each new codon in the mRNA being
translated, resulting in the synthesis of a polypeptide chain that may be several hundred
residues long. Eventually, the translation complex reaches the final codon at the end of
the coding region, where translation is terminated.

The elongation reactions in eukaryotes are very similar to those in E. coli Three ac-
cessory protein factors participate in chain elongation in eukaryotes: EF-1 a, EF-1/3,
and EF-2. EF-1 a docks the aminoacyl-tRNA in the A site; its activity thus parallels that
of E. coli EF-Tu. EF-1 (3 acts like bacterial EF-Ts, recycling EF-la. EF-2 carries out
translocation in eukaryotes. EF-Tu and EF-1 a are highly conserved, homologous pro-
teins, as are EF-G and EF-2. Eukaryotic and prokaryotic ribosomal RNAs are also very
similar in sequence and in secondary structure. These similarities indicate that the com-
mon ancestor of prokaryotes and eukaryotes carried out protein synthesis in a manner
similar to that seen in modern organisms. Thus, protein synthesis is one of the most an-
cient and fundamental biochemical reactions.

22.7 Termination of Translation

E. coli has three release factors (RF-1, RF-2, and RF-3) that participate in the termina-
tion of protein synthesis. After formation of the final peptide bond, the peptidyl-tRNA
is translocated from the A site to the P site, as usual. The translocation positions one of
the three termination codons (UGA, UAG, or UAA) in the A site. These termination
codons are not recognized by any tRNA molecules so protein synthesis stalls at the
termination codon. Eventually, one of the release factors diffuses into the A site. RF-1
recognizes UAA and UAG and RF-2 recognizes UAA and UGA. RF-3 binds GTP and
enhances the effects of RF-1 and RF-2.

When the release factors recognize a termination codon, they cause hydrolysis of
the peptidyl-tRNA. Release of the completed polypeptide is probably accompanied by
GTP hydrolysis and dissociation of the release factors from the ribosome. At this point,
the ribosomal subunits dissociate from the mRNA and initiation factors bind to the 30S
subunit in preparation for the next round of protein synthesis.

22.8 Protein Synthesis Is Energetically
Expensive

Protein synthesis is very expensive — it uses a large fraction of all ATP equivalents that
are available in a cell. Where does all this energy go?

For each amino acid added to a polypeptide chain, four phosphoanhydride
bonds are cleaved: ATP is hydrolyzed to AMP + 2 Pi during activation of the amino
acid and two GTP molecules are hydrolyzed to 2 GDP + 2 Pi during chain elonga-
tion. The hydrolysis of GTP is coupled to conformational changes in the translation
machinery. In this sense, GTP and GDP act as allosteric modulators. However, unlike
most conformational changes induced by allosteric modulators, the conformational
changes that occur during protein synthesis are associated with a considerable con-
sumption of energy.

The hydrolysis of four phosphoanhydride bonds represents a large Gibbs free en-
ergy change — much more than is required for the formation of a single peptide bond.
Most of the “extra” energy compensates for the loss of entropy during protein synthesis.
The decrease in entropy is due primarily to the specific ordering of 20 different kinds of
amino acids into a polypeptide chain. In addition, entropy is lost when an amino acid is
linked to a particular tRNA and when an aminoacyl-tRNA associates with a specific
codon.

22.9 Regulation of Protein Synthesis 685

22.9 Regulation of Protein Synthesis

One way gene expression can be regulated is by controlling the translation of mRNA
into protein. Translation can be controlled at initiation, elongation, or termination. In
general, translational control of gene expression is used to regulate the production of
proteins that assemble into multisubunit complexes and proteins whose expression in
the cell must be strictly and quickly controlled.

The rate of translation depends to some extent on the sequence of the template.
An mRNA containing an abundance of rare codons, for example, is translated less rap-
idly (and therefore less frequently) than one containing the most frequently used codons.
In addition, the rate of translation initiation varies with the nucleotide sequence at the
initiation site. A strong ribosome binding site in bacterial mRNA leads to more efficient
initiation. There is also evidence that the nucleotide sequence surrounding the initiation
codon in eukaryotic mRNA influences the rate of initiation.

One difference between the initiation of translation and the initiation of transcrip-
tion is that the formation of a translation complex can be influenced by secondary
structure in the message. For example, the formation of intramolecular double-
stranded regions in mRNA can mask ribosome binding sites and the initiation codon.
Although structural properties can determine whether a given mRNA molecule is
translated frequently or infrequently, this is not regulation in the strict sense. We use the
term translational regulation to refer to cases where extrinsic factors modulate the fre-
quency of mRNA translation.

Ribosomes moving
on messenger RNA
synthesize proteins
haiku by Sydney Brenner (2002)

Polypeptide synthesis is an example
of head growth (Box 1 2.5).

KEY CONCEPT

mRNA codons in the ribosome’s A site are
also being continually tested by randomly
diffusing release factors, which are
seeking translation termination codons.

A. Ribosomal Protein Synthesis Is Coupled to Ribosome
Assembly in f. coli

Every E. coli ribosome contains at least 52 ribosomal proteins. The genes encoding these
ribosomal proteins are scattered throughout the genome in 13 operons and seven iso-
lated genes. When multiple copies of genes encoding some of these ribosomal proteins
are inserted into E. coli , the concentrations of the respective mRNAs increase sharply,
yet the overall rate of ribosomal protein synthesis scarcely changes. Furthermore, the
relative concentrations of ribosomal proteins remain unchanged even though the vari-
ous mRNA molecules for ribosomal proteins are present in unequal amounts. These
findings suggest that the synthesis of ribosomal proteins is tightly regulated at the level
of translation.

Translational regulation of ribosomal protein synthesis is crucial since ribosomes
cannot assemble unless all the proteins are present in the proper stoichiometry. The
production of ribosomal proteins is controlled by regulating the efficiency with which
their mRNAs are translated. Each of the large operons containing ribosomal protein
genes encodes one ribosomal protein that inhibits translation of its own polycistronic
mRNA by binding near the initiation codon of one of the first genes of the operon.

The interactions between the inhibiting ribosomal proteins and their mRNAs may
resemble the interactions between these proteins and the ribosomal RNA to which they
bind when assembled into mature ribosomes. For example, the mRNA transcript of the
str operon, which includes the coding region for the ribosomal protein S7, contains some
regions of RNA sequence that are identical to the S7 binding site of 16S rRNA. Moreover,
the proposed secondary structure of the str mRNA resembles the proposed secondary
structure of the 16S rRNA S7 binding site (Figure 22.25). S7 binds to this region of the str
mRNA molecule and inhibits translation. It is likely that S7 recognizes analogous struc-
tural features in both RNA molecules. Similar mechanisms regulate the translation of
mRNAs that encode the other ribosomal proteins.

The ribosomal proteins that inhibit translation bind more tightly to ribosomal
RNA than to the similar sites on messenger RNA. Thus, the mRNA continues to be
translated as long as newly synthesized ribosomal proteins are incorporated into ribo-
somes. However, as soon as ribosome assembly slows and the concentration of free
ribosomal proteins increases within the cell, the inhibiting ribosomal proteins bind to
their own mRNA molecules and block additional protein synthesis. In this way, synthe-
sis of ribosomal proteins is coordinated with ribosome assembly.

686 CHAPTER 22 Protein Synthesis

BOX 22.1 SOME ANTIBIOTICS INHIBIT PROTEIN SYNTHESIS

Many microorganisms produce antibiotics, which they use as
a chemical defense against competitors. Some antibiotics pre-
vent bacterial growth by inhibiting the formation of peptide
bonds. For example, the structure of the antibiotic puromycin
closely resembles the structure of the 3' end of an aminoacyl-
tRNA molecule. Because of this similarity, puromycin can
enter the A site of a ribosome. Peptidyl transferase then cat-
alyzes the transfer of the nascent polypeptide to the free
amino group of puromycin (see figure below). The peptidyl -
puromycin is bound weakly in the A site and soon dissociates
from the ribosome, thereby terminating protein synthesis.

Although puromycin effectively blocks protein synthesis
in prokaryotes, it is not clinically useful since it also blocks

protein synthesis in eukaryotes and is therefore poisonous to
humans. Clinically important antibiotics, which include
streptomycin, chloramphenicol, erythromycin, and tetracy-
cline, are specific for bacteria and have little or no effect on
eukaryotic protein synthesis. Streptomycin binds to one of
the ribosomal proteins in the 30S subunit and inhibits the
initiation of translation. Chloramphenicol interacts with the
50S subunit and inhibits peptidyl transferase. Erythromycin
binds to the 50S subunit, inhibiting the translocation step.
Tetracycline binds to the 3 OS subunit, preventing the binding
of aminoacyl-tRNA molecules to the A site.

tRNA

▲ Formation of a peptide bond between puromycin at the A site of a ribosome and the nascent
peptide bound to the tRNA in the P site. The product of this reaction is bound only weakly in the
A site and dissociates from the ribosome, thus terminating protein synthesis and producing an
incomplete, inactive peptide.

B. Globin Synthesis Depends on Heme Availability

The synthesis of hemoglobin, the major protein in red blood cells, requires globin
chains and heme in stoichiometric amounts (Section 4.12). One way globin synthesis
is controlled is by regulation of translation initiation. Hemoglobin is initially synthe-
sized in immature erythrocytes called rubriblasts. Mammalian rubriblasts lose
their nuclei during maturation and eventually become reticulocytes, which are the

22.9 Regulation of Protein Synthesis 687

immediate precursors of erythrocytes. Hemoglobin continues to be synthesized in
reticulocytes that are packed with processed, stable mRNA molecules encoding glo-
bin polypeptides.

The rate of globin synthesis in reticulocytes is determined by the concentration
of heme. When the concentration of heme decreases, the translation of globin mRNA
is inhibited. The effect of heme on globin mRNA translation is mediated by a protein
kinase called heme-controlled inhibitor (HCI) (Figure 22.26). Active HCI catalyzes
transfer of a phosphoryl group from ATP to the translation initiation factor eIF-2.
Phosphorylated eIF-2 is unable to participate in translation initiation and protein
synthesis in the cell is inhibited.

During the initiation of translation, eIF-2 binds methionyl-tRNA^ 61 and GTP.
When the preinitiation complex encounters an initiation codon, methionyl-
tRNAi Met is transferred from eIF-2 to the initiation codon of the mRNA. This
transfer reaction is accompanied by the hydrolysis of GTP and the release of elF-
2-GDP. An enzyme called guanine nucleotide exchange factor (GEF) catalyzes the
replacement of GDP with GTP on eIF-2 and the attachment of another
methionyl-tRNAi Met to eIF-2. GEF binds very tightly to phosphorylated elF-
2-GDP, preventing the nucleotide exchange reaction. Protein synthesis is com-
pletely inhibited when all the GEF in the cell is bound because the active elF-
2-GTP complex cannot be regenerated.

Heme regulates the synthesis of globin by interfering with the activation of
HCI. When heme is abundant, HCI is inactive and globin mRNA can be trans-
lated. When heme is scarce, however, HCI is activated and translation of all mRNA
within the cell is inhibited (Figure 22.26). Phosphorylation of eIF-2 also appears to
regulate the translation of mRNA in other mammalian cell types. For example,
during infection of human cells by RNA viruses, the presence of double-stranded
RNA leads to the production of interferon, which in turn activates a protein kinase
that phosphorylates eIF-2. This reaction inhibits protein synthesis in the virus-
infected cell.

C. The f. coli trp Operon Is Regulated by Repression and Attenuation

The trp operon in E. coli encodes the proteins necessary for the biosynthesis of trypto-
phan. Most organisms synthesize their own amino acids but can also obtain them by
degrading exogenous proteins. For this reason, most organisms have evolved mechanisms

▲ Figure 22.25

Comparison of proposed secondary structures
of S7 binding sites, (a) S7 binding site on
16S rRNA. (b) S7 binding site on the str
mRNA molecule.

and unable to catalyze
nucleotide exchange
for eIF-2.

◄ Figure 22.26

Inhibition of protein synthesis by phosphoryla-
tion of eIF-2 in reticulocytes. When the con-
centration of heme is high, HCI is inactive
and translation proceeds normally. When the
concentration of heme is low, HCI catalyzes
the phosphorylation of eIF-2. Phosphory-
lated eIF-2 binds the limiting amounts of
GEF in the cell very tightly, sequestering the
GEF and preventing translation of cellular
mRNAs (including the globins).

688

CHAPTER 22 Protein Synthesis

t Figure 22.27

Repression of the E. coli trp operon. The trp

operon is composed of a leader region and
five genes required for the biosynthesis of
tryptophan from chorismate. The trp R gene,
located upstream of the trp operon ( trpO ),
encodes trp repressor, which is inactive in
the absence of its corepressor, tryptophan.
When tryptophan is present in excess, it
binds to trp repressor, and the repressor-
tryptophan complex binds to the trp operator
( trpO ). Once bound to the operator, the
repressor-tryptophan complex prevents
further transcription of the trp operon by ex-
cluding RNA polymerase from the promoter.

to repress the synthesis of the enzymes required for de novo amino acid biosynthesis
when the amino acid is available from exogenous sources. For example, in E. coli, tryp-
tophan is a negative regulator of its own biosynthesis. In the presence of tryptophan,
the trp operon is not expressed (Figure 22.27). Expression of the trp operon is inhib-
ited in part by trp repressor, a dimer of two identical subunits, trp repressor is en-
coded by the trpR gene, which is located elsewhere on the bacterial chromosome and is
transcribed separately. When tryptophan is abundant, a repressor- tryptophan complex
binds to the operator trpO , which lies within the promoter. The bound repressor-
tryptophan complex prevents RNA polymerase from binding to the promoter. Trypto-
phan is thus a corepressor of the trp operon.

Regulation of the E. coli trp operon is supplemented and refined by a second, in-
dependent mechanism called attenuation. This second mechanism depends on transla-
tion and helps determine whether transcription of the trp operon proceeds or termi-
nates prematurely. The movement of RNA polymerase from the promoter into the
trpE gene is governed by a 162 nucleotide sequence that lies between the promoter
and trpE. This sequence, called the leader region (Figure 22.27), includes a stretch of
45 nucleotides that encodes a 14 amino acid peptide called the leader peptide. The
mRNA transcript of the leader region contains two consecutive codons specifying
tryptophan near the end of the coding region for the leader peptide. In addition, the

Promoter

22.10 Post-Translational Processing

689

leader region contains four GC-rich sequences. The codons that specify tryptophan
and the four GC-rich sequences regulate the synthesis of mRNA by affecting tran-
scription termination.

When transcribed into mRNA, the four GC-rich sequences of the leader region can
base-pair to form one of two alternative secondary structures (Figure 22.28, on the next
page). The first possible secondary structure includes two RNA hairpins. These hairpins
form between the sequences labeled 1 and 2 and between those labeled 3 and 4 in Figure
22.28a. The 1-2 hairpin is a typical transcription pause site. The 3-4 hairpin is followed by a
string of uridylate residues, which is a typical rho- independent termination signal (Section
21.4). This particular termination signal is unusual, however, because it occurs upstream
of the first gene in the trp operon. The other possible secondary structure includes a
single RNA hairpin between sequences 2 and 3. This hairpin, which is more stable than
the 3-4 hairpin, forms only when sequence 1 is not available for hairpin formation with
sequence 2.

During transcription of the leader region, RNA polymerase pauses when the 1-2
hairpin forms. While RNA polymerase pauses, a ribosome initiates translation of the
mRNA encoding the leader peptide. This coding region begins just upstream of the 1-2
RNA hairpin. Sequence 1 encodes the C-terminal amino acids of the leader peptide and
also contains a termination codon. As the ribosome translates sequence 1, it disrupts the
1-2 hairpin, thereby releasing the paused RNA polymerase, which then transcribes se-
quence 3. In the presence of tryptophanyl-tRNA Trp , the ribosome and RNA polymerase
move at about the same rate. When the ribosome encounters the termination codon of
the trp leader mRNA, it dissociates and the 1-2 hairpin re-forms. After the ribosome has
disassembled, RNA polymerase transcribes sequence 4, which forms a transcription ter-
mination hairpin with sequence 3. This termination signal causes the transcription
complex to dissociate from the DNA template before the genes of the trp operon have
been transcribed.

When tryptophan is scarce, however, the ribosome and RNA polymerase do not
move synchronously When the concentration of cellular tryptophan falls, the cell be-
comes deficient in tryptophanyl-tRNA Trp . Under these circumstances, the ribosome
pauses when it reaches the two codons specifying tryptophan in sequence 1 of the
mRNA molecule. RNA polymerase, which has already been released from the 1-2
pause site, transcribes sequences 3 and 4. While the ribosome is stalled and sequence 1
is covered, sequence 2 forms a hairpin loop with sequence 3. Since the 2-3 hairpin is
more stable than the 3-4 hairpin, sequence 3 does not pair with sequence 4 to form the
transcription termination hairpin. Under these conditions, RNA polymerase passes
through the potential termination site (UGA in Figure 22.28a), and the rest of the trp
operon is transcribed.

Attenuation appears to be a regulatory mechanism that has evolved relatively re-
cently and is found only in enteric bacteria, such as E. coli. (Attenuation cannot
occur in eukaryotes because transcription and translation take place in different
parts of the cell.) Several E. coli operons, including the phe, thr, his, leu, and He oper-
ons, are regulated by attenuation. Some operons, such as the trp operon, combine at-
tenuation with repression, whereas others, such as the his operon, are regulated
solely by attenuation. The leader peptides of operons whose genes are involved in
amino acid biosynthesis may contain as many as seven codons specifying a particular
amino acid.

22.10 Post-Translational Processing

As the translation complex moves along the mRNA template in the 5' —> 3' direction,
the nascent polypeptide chain lengthens. The 30 or so most recently polymerized
amino acid residues remain buried in the ribosome, but amino acid residues closer to
the N- terminus are extruded from the ribosome. The N- terminal residues start to
fold into the native protein structure even before the C-terminus of the protein has

690 CHAPTER 22 Protein Synthesis

(a) Leader peptide ^

Met Lys Ala lie Phe Val Leu Lys Gly Trp Trp Arg Thr Ser STOP
AaTg AAA GOA AAHJ uTTc (TuA ClTg AAA cTgU iTgG iTgG (TgC AcTj iTcC lTgI

A 30 40 50 60 70

GCUAUGGGAAA.

20 A

70 A

2 c A

GCGUACCACUUAUGUGACGGG C

pppAAG U U C AC G 10
5'

^AGCAAUCAGAUACCCAGCCCGCCU,

100 110 120 A

4 1 A

r AUUAAAACAAGUUUUUUUUCGGGCGAG U

i ° 150 140 130

Met Gin Thr

vA/VW'

UAACAAUGCAAACA

160

trpE Polypeptide

WVAA.

(b)

vAAAA/' A

STOP

vAA/VV'A

2-3 Hairpin
A

140

u u u

v/VWV'

U
G
C-
G-
U— A
A A

- G ioo

-C

90 C

c —

u —

U A-

u —

G —

80 G

C —

G —

C —

140

U U U

JWW

1-2

Transcription

pause structure
M 3-4

Transcription

termination

signal

▲ Figure 22.28

trp leader region, (a) mRNA transcript of the trp leader region. This 162 nucleotide mRNA sequence includes four GC-rich sequences and the coding
region for a 14 amino acid leader peptide. The coding region includes two consecutive codons specifying tryptophan. The four GC-rich sequences
can base-pair to form one of two alternative secondary structures, (b) Sequence 1 (red) and sequence 2 (blue) are complementary and, when base-
paired, form a typical transcription pause site. Sequence 3 (green) and sequence 4 (yellow) are complementary and, when base-paired, form a rho-
independent termination site, (c) Sequences 2 and 3 are also complementary and can form an RNA hairpin that is more stable than the 3-4 hairpin.
This structure forms only when sequence 1 is not available for hairpin formation with sequence 2.

been synthesized. As these residues fold, they are acted on by enzymes that modify the
nascent chain.

Modifications that occur before the polypeptide chain is complete are said to be
cotranslational, whereas those that occur after the chain is complete are said to be post-
translational. Some examples from the multitude of cotranslational and post-transla-
tional modifications include deformylation of the N-terminal residue in prokaryotic

22.10 Post-Translational Processing 691

proteins, removal of the N-terminal methionine from prokaryotic
and eukaryotic proteins, formation of disulfide bonds, cleavage by
proteinases, phosphorylation, addition of carbohydrate residues,
and acetylation.

One of the most important events that occurs co- and post-
translationally is the processing and transport of proteins through
membranes. Protein synthesis occurs in the cytosol, but the ma-
ture forms of many proteins are embedded in membranes or are
inside membrane bounded compartments. For example, many re-
ceptor proteins are embedded in the external membrane of the
cell, with the bulk of the protein outside the cell. Other proteins
are secreted from cells, and still others reside in lysosomes and
other organelles inside eukaryotic cells. In each case, the protein
synthesized in the cytosol must be transported across a membrane
barrier. In fact, such proteins are synthesized by membrane bound
ribosomes that are attached to the plasma membrane in bacteria
and to the endoplasmic reticulum in eukaryotic cells.

The best-characterized transport system is the one that car-
ries proteins from the cytosol to the plasma membrane for secretion
(Figure 22.29). In eukaryotes, proteins destined for secretion are
transported across the membrane of the endoplasmic reticulum
into the lumen, which is topologically equivalent to the cell exte-
rior. Once the protein has been transported into the endoplasmic
reticulum, it can be transported by vesicles through the Golgi
apparatus to the plasma membrane for release outside the cell.

A. The Signal Hypothesis

Cytosol

Lumen of
endoplasmic
reticulum

Secreted proteins are synthesized on the surface of the endoplas-
mic reticulum, and the newly synthesized protein is passed
through the membrane into the lumen. In cells that make large
amounts of secreted protein, the endoplasmic reticulum mem-
branes are covered with ribosomes (Figure 22.30, on the next page).

The clue to the process by which many proteins cross the
membrane of the endoplasmic reticulum appears in the first 20 or
so residues of the nascent polypeptide chain. In most membrane
bound and secreted proteins, these residues are present only in the
nascent polypeptide, not in the mature protein. The N-terminal
sequence of residues that is proteolytically removed from the pro-
tein precursor is called the signal peptide since it is the portion of
the precursor that signals the protein to cross a membrane. Signal
peptides vary in length and composition, but they are typically
from 16 to 30 residues long and include 4 to 15 hydrophobic
residues (Figure 22.31, on the next page).

In eukaryotes, many proteins destined for secretion appear to be translocated across
the endoplasmic reticulum by the pathway shown in Figure 22.32 on page 693. In the
first step, an 80S initiation complex — including a ribosome, a Met-tRNAi Met molecule,
and an mRNA molecule — forms in the cytosol. Next, the ribosome begins translating the
mRNA and synthesizing the signal peptide at the N- terminus of the precursor. Once the
signal peptide has been synthesized and extruded from the ribosome, it binds to a pro-
tein-RNA complex called a signal recognition particle (SRP).

SRP is a small ribonucleoprotein containing a 300 nucleotide RNA molecule called
7SL RNA and four proteins. SRP recognizes and binds to the signal peptide as it
emerges from the ribosome. When SRP binds, further translation is blocked. The SRP-
ribosome complex then binds to an SRP receptor protein (also known as docking pro-
tein) on the cytosolic face of the endoplasmic reticulum. The ribosome is anchored to
the membrane of the endoplasmic reticulum by ribosome binding proteins called
translocons, and the signal peptide is inserted into the membrane at a pore that is part

Extracellular

space

Plasma

membrane

▲ Figure 22.29

Secretory pathway in eukaryotic cells.

Proteins whose synthesis begins in the cy-
tosol are transported into the lumen of the
endoplasmic reticulum. After further modifi-
cation in the Golgi apparatus, the proteins
are secreted.

692 CHAPTER 22 Protein Synthesis

Figure 22.30 ►

Secretory vesicles in a maize rootcap cell.

Large secretory vesicles containing proteins
are budding off the Golgi apparatus
(center). Note the abundance of ribosomes
bound to the endoplasmic reticulum.

Golgi apparatus

Mitochondrion '

E nd op I as m i c , ret i cu I u m

1 Vesicles

A ^

Ribosomes

Golgi apparatus

of the complex formed by the endoplasmic reticulum proteins at the docking site. Once
the ribosome-SRP complex is bound to the membrane, the inhibition of translation is
relieved and SRP dissociates in a reaction coupled to GTP hydrolysis. Thus, the role of
SRP is to recognize nascent polypeptides containing a signal peptide and to target the
translation complex to the surface of the endoplasmic reticulum.

Once the translation complex is bound to the membrane, translation resumes and
the new polypeptide chain passes through the membrane. The signal peptide is then
cleaved from the nascent polypeptide by a signal peptidase, an integral membrane pro-
tein associated with the pore complex. The transport of proteins across the membrane

Prelysozyme

® i

H 3 N-Met-Arg-Ser-Lei -Leu- - Bu-Val-Leu-Cys-Phe-Leu-Pn - ?u-Ala-Ala-Leu-Gly-Gly

Preproalbumin

H 3 N-Met-Lys-Trp-Val-Thr-Phe-Leu-Leu-Leu- iu-Phc- -Ser-Gly-Ser-AI - le-Ser-Arg^x^

Alkaline phosphatase

H 3 N-Met-Lys-Gln-Ser-Thr- le-Ala-Le - la-Leu-Lei -Pro-Leu- ei -Phe-Thr-Pro-Va -Thr-Lys-Ala-Arg

Maltose-binding protein

H 3 N-Met-Lys-i e-Lys-Thr-Gly- da-Arg- -Le - la-Leu-Ser-Ala- -Thr-Thr-Met-Met-Phe-Ser-Ala-Ser-AI; -Leu-Ala-Lys^^
OmpA

H 3 N-Met-Lys-Lys-Thr-Ala-lle-Ala-lle-Ala-Val-Ala-Leu-Ala-Gly-Phe-Ala-Thr-Val-Ala-Gln-Ala-Ala

Figure 22.31 ▼

Signal peptides from secreted proteins.

Hydrophobic residues are shown in blue,
and arrows mark the sites where the signal
peptide is cleaved from the precursor.
(OmpA is a bacterial membrane protein.)

22.10 Post-Translational Processing 693

SRP

mRNA

Signal

peptide

SRP binds to the signal
peptide as it emerges from
the ribosome. Translation
is inhibited.

Plasma membrane
of endoplasmic
reticulum

SRP binds to the SRP
receptor on the surface of
the endoplasmic reticulum.

Translocon

SRP
receptor

GTP

peptidase

GDP

The ribosome binds to
translocon, and the signal
peptide is inserted through
a pore in the membrane.
Translation resumes.

◄ Figure 22.32

Translocation of eukaryotic proteins into the
lumen of the endoplasmic reticulum.

Subsequent translation passes
the nascent polypeptide into
the lumen of the endoplasmic
reticulum. The signal peptide
is removed by signal peptidase

694 CHAPTER 22 Protein Synthesis

Figure 22.33 ►

Structure of a complex oligosaccharide linked
to an asparagine residue. Abbreviations: Glc,
glucose; GIcNAc, /V-acetylglucosamine;

Man, mannose.

is assisted by chaperones in the lumen of the endoplasmic reticulum. In addition to
their role in protein folding, chaperones are required for translocation, and their activity
requires ATP hydrolysis. When protein synthesis terminates, the ribosome dissociates
from the endoplasmic reticulum, and the translation complex disassembles.

B. Glycosylation of Proteins

Many integral membrane proteins and secretory proteins contain covalently bound
oligosaccharide chains. The addition of these chains to proteins is called protein glyco-
sylation (Section 8.7C). Protein glycosylation is one of the major metabolic activities of
the lumen of the endoplasmic reticulum and of the Golgi apparatus and is an extension
of the general process of protein biosynthesis. A glycoprotein can contain dozens, in-
deed hundreds, of monosaccharide units. The mass of the carbohydrate portion may
account for as little as 1% or as much as 80% of the mass of the glycoprotein.

A common glycosylation reaction involves the covalent attachment of a complex
oligosaccharide to the side chain of an asparagine residue (Figure 22.33). During subse-
quent transit through the endoplasmic reticulum and the Golgi apparatus, proteins
may be covalently modified in many ways, including the formation of disulfide bonds
and proteolytic cleavage. The complex oligosaccharides attached to the proteins are
likewise modified during transit. A variety of different oligosaccharides can be cova-
lently bound to proteins. In some cases, the structure of the oligosaccharide acts as a
signal to target proteins to a specific location. For example, lysosomal proteins contain
sites for the attachment of an oligosaccharide that targets these proteins to the lyso-
some. By the time they have traversed the Golgi apparatus, the proteins and their
oligosaccharides are usually fully modified.

Summary

1. The genetic code consists of nonoverlapping, three-nucleotide
codons. The code is unambiguous and degenerate; the first two
nucleotides of the three-letter code are often sufficient; codons
with similar sequences specify chemically similar amino acids;
and there are special codons for the initiation and termination of
peptide synthesis.

2. tRNA molecules are the adapters between mRNA codons and
amino acids in proteins. All tRNA molecules have a similar
cloverleaf secondary structure with a stem and three arms. The
tertiary structure is L-shaped. The anticodon loop is at one end of
the structure, and the acceptor stem is at the other. The anticodon
in tRNA base-pairs with a codon in mRNA. The 5' (wobble) posi-
tion of the anticodon is conformationally flexible.

3. An aminoacyl-tRNA synthetase catalyzes the addition of a spe-
cific amino acid to the acceptor stem of the appropriate tRNA,
producing an aminoacyl-tRNA. Some aminoacyl-tRNA syn-
thetases carry out proofreading.

4. Ribosomes are the RNA-protein complexes that catalyze the poly-
merization of amino acids bound to aminoacyl-tRNA molecules.
All ribosomes are composed of two subunits: prokaryotic ribo-
somes contain three rRNA molecules, and eukaryotic ribosomes
contain four. The growing polypeptide chain is attached to a
tRNA in the peptidyl (P) site of the ribosome, and the aminoacyl-
tRNA molecule bearing the next amino acid to be added to the
nascent polypeptide chain docks in the aminoacyl (A) site.

5. Translation begins with the formation of an initiation complex
consisting of an initiator tRNA, the mRNA template, the ribosomal
subunits, and several initiation factors. In prokaryotes, initiation
occurs just downstream of Shine-Dalgarno sequences; in eukary-
otes, initiation usually occurs at the initiation codon closest to the
5' end of the mRNA.

6. The elongation step of translation requires accessory proteins
called elongation factors. The three steps of elongation are
(1) positioning of the correct aminoacyl-RNA in the A site,

Problems 695

(2) formation of the peptide bond by peptidyl transferase, and

(3) translocation of the ribosome by one codon.

7. Release factors recognize termination codons and catalyze the ter-
mination of protein synthesis and disassembly of the translation
complex.

8. Protein synthesis requires the energy of four phosphoanhydride
bonds per residue.

9. The regulation of translation includes the formation of secondary
structure in mRNA that influences the rate of initiation. Riboso-

mal RNA proteins can inhibit translation of their own mRNA by
binding to such sites. Phosphorylation of an initiation factor reg-
ulates globin synthesis. Regulation of expression of the E. coli trp
operon involves attenuation, in which translation of a leader
mRNA governs transcription of the operon.

10 . Many proteins are post-translationally modified. Some eukary-
otic proteins destined for secretion contain N-terminal signals for
transport into the endoplasmic reticulum. Many membrane and
secreted proteins are glycosylated.

Problems

1. The standard genetic code is read in codons that are three nu-
cleotides long. How many potential reading frames are there on a
single piece of double-stranded DNA? If instead the genetic code
was read in codons that were four nucleotides long, how many
reading frames would there be on the same piece of double-
stranded DNA?

2. Examine the sequences of the mRNAs transcribed from the DNA
sequence in Problem 1 1 in Chapter 21. Assuming that the DNA
segment is from the middle of a protein-coding gene, which of
the possible mRNAs is most likely to be the actual transcript?
What is the sequence of the encoded peptide?

3. Calculate the number of phosphoanhydride bonds that are hy-
drolyzed during synthesis of a 600 amino acid residue protein in
E. coli. Do not include the energy required to synthesize the
amino acids, mRNA, tRNA, or the ribosomes.

4. Polypeptide chain elongation on the ribosome can be broken
down into three discrete steps (the microcycle): (1) binding of the
correct aminoacyl-tRNA in the ribosome’s A site, (2) peptide
bond formation, and (3) translocation. What, specifically, is it
that gets translocated in the third step of this cycle?

5. A prokaryotic mRNA may contain many AUG codons. How does
the ribosome distinguish AUG codons specifying initiation from
AUG codons specifying internal methionine?

6. Given that the genetic code is universal, would a plant mRNA be
correctly translated in a prokaryotic cell like E. coli ?

7. Bacterial genomes usually contain multiple copies of the genes for
rRNA. These are transcribed very efficiently in order to produce
large amounts of rRNA for assembly into ribosomes. In contrast,
the genes that encode ribosomal proteins are present only as sin-
gle copies. Explain the difference in the number of copies of
rRNA and ribosomal protein genes.

8. Suppressor mutations suppress the effects of other mutations. For
example, mutations that produce the stop codon UAG in the mid-
dle of a gene are suppressed by an additional mutation in a tRNA
gene that gives rise to a mutant anticodon with the sequence
CUA. Consequently, an amino acid is inserted at the mutant stop
codon, and a protein is synthesized (although it may be only par-
tially active). List all the tRNA species that could be mutated to a
suppressor of UAG mutations by a single base change in the anti-
codon. How can a cell with a suppressor tRNA survive?

9. Transfer RNAs are absolutely essential for polypeptide synthesis.
After reviewing the material in this chapter, name five different cel-
lular components that can bind to (interact with) tRNA molecules.

10 . On rare occasions, the translation machinery encounters a codon
that cannot be quickly interpreted because of the lack of a partic-
ular tRNA or release factor. In these cases, the ribosome may
pause and then shift by a single nucleotide and begin translating a
different reading frame. Such an occurrence is known as transla-
tional frameshifting. The E. coli release factor RF-2, which is
translated from mRNA that contains an internal UGA stop
codon, is produced by translational frameshifting. Explain how
this phenomenon might regulate RF-2 production.

11. The mechanism of attenuation requires the presence of a leader
region. Predict the effect of the following changes on regulation
of the trp operon:

(a) The entire leader region is deleted.

(b) The sequence encoding the leader peptide is deleted.

12 . In Chapter 21, you learned of many different regulatory mecha-
nisms that control transcription of the lac operon in E. coli. In
Chapter 22, one of the mechanisms of translational regulation
discussed was called attenuation. Would you predict that in some
other bacterial species the lac operon might have evolved such
that an attenuation mechanism was used to regulate expression
levels from this operon?

13. In the operons that contain genes for isoleucine biosynthesis, the
leader regions that precede the genes contain multiple codons
that specify not only isoleucine but valine and leucine as well.
Suggest a reason why this is so.

14 . Suggest the steps involved in the synthesis and processing of a gly-
cosylated, eukaryotic integral membrane protein with a C-termi-
nal cytosolic domain and an N-terminal extracellular domain.

15 . In Chapter 23, you will learn about recombinant DNA techniques
that allow genes to be cut and pasted at will. If you could remove
the coding region for a secretion signal sequence from one pro-
tein and place it such that it will now occupy the N-terminus of a
cytosolic protein (e.g., /3-galactosidase), would you expect the
new hybrid protein to enter the cell’s secretory pathway?

16 . In some species of bacteria, the codon GUG initiates protein syn-
thesis (e.g., LacI, Figure 22.17a). The completed proteins always
contain methionine at the N-terminus. How can the initiator
tRNA base-pair with the codon GUG? How is this phenomenon
related to wobble?

696 CHAPTER 22 Protein Synthesis

Selected Readings

Aminoacyl-tRNA Synthetases

Carter, C. W., Jr. (1993). Cognition, mechanism,
and evolutionary relationships in aminoacyl-tRNA
synthetases. Annu. Rev. Biochem. 62:715-748.

Ibba, M., and Soil, D. (2000). Aminoacyl-tRNA
synthesis. Annu. Rev. Biochem. 69:617-650.

Jakubowski, H., and Goldman, E. (1992). Editing
of errors in selection of amino acids for protein
synthesis. Microbiol. Rev. 56:412-429.

Kurland, C. G. (1992). Translational accuracy and
the fitness of bacteria. Annu. Rev. Genet. 26:29-50.

Schimmel, R, and Ribas de Pouplana, L. (2000).
Footprints of aminoacyl-tRNA synthetases are
everywhere. Trends Biochem. Sci. 25:207-209.

Ribosomes and Translation

Ban, N., Nissen, R, Hansen, J., Moore, P. B., and
Steitz, T. A. (2000). The complete atomic structure
of the large ribosomal subunit at 2.4A resolution.
Science 289:905-919.

Carter, A. P., Clemons, W. M., Brodersen, D. E.,
Morgan-Warren, R. J., Wimberly, B. T., and
Ramakrishnan, V. (2000). Functional insights from
the structure of the 30S ribosomal subunit and its
interactions with antibiotics. Nature 407:340-348.

Garrett, R. A., Douthwate, S. R., Matheson A. T.,
Moore, P. B., and Noller, H. F., eds. (2000). The
Ribosome: Structure, Function, Antibiotics and
Cellular Interactions (Washington, DC: American
Society for Microbiology).

Hanawa-Suetsugu, K., Sekine, S., Sakai, H., Hori-
Takemoto, C., Tevader, T., Unzai, S., Tame, J.R.H.,
Kuramitsu, S., Shirouzu, M., and Yokoyama, S.
(2004). Crystal structure of elongation factor P
from Thermus thermophilus HB8. Proc. Natl. Acad.
Sci. 101:9595-9600.

Kawashima, T., Berthet-Colominas, C., Wulff, M.,
Cusack, S., and Leberman, R. (1996). The structure
of the Escherichia coli EF-Tu • EF-Ts complex at
2.5 A resolution. Nature 379:51 1-518.

Moore, P. B., and Steitz, T. A. (2003). The struc-
tural basis of large ribosomal subunit function.
Annu. Rev. Biochem. 72:813-850.

Nirenberg, M.W., and Matthaei, J.H., (1961). The
dependence of cell-free protein synthesis in E. coli
upon naturally occurring or synthetic polyribo-
nucleotides. Proc. Natl. Acad. Sci. 47:1588-1602.

Noller, H. F. (1993). Peptidyl transferase: protein,
ribonucleoprotein, or RNA? /. Bacteriol.
175:5297-5300.

Pestova, T. V., and Hellen, C. U. T. (1999). Ribo-
some recruitment and scanning: what’s new?
Trends Biochem. Sci. 24:85-8 7.

Ramakrishnan, V. (2009). Unravelling the struc-
ture of the ribosome. Nobel Fecture 135-160.

Selmer, M., Al-Karadaghi, S., Hirokawa,

G., Kaji, A., and Filjas, A. (1999). Crystal Structure
of Thermotoga maritima ribosome recycling factor:
A tRNA mimic. Science 286:2349-2352.

Steitz, T.A. (2009). From the structure and function
of the ribosome to new antibiotics. Nobel Fecture
179-204.

Regulation of Translation

Kozak, M. (1992). Regulation of translation in eu-
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McCarthy, J. E. G., and Gualerzi, C. (1990). Trans-
lational control of prokaryotic gene expression.
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Merrick, W. C. (1992). Mechanism and regulation
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56:291-315.

Rhoads, R. E. (1993). Regulation of eukaryotic pro-
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268:3017-3020.

Samuel, C. E. (1993). The eIF-2a protein kinases,
regulators of translation in eukaryotes from yeasts
to humans. /. Biol. Chem. 268:7603-7606.

Post-translational Modification

Hurtley, S. M. (1993). Hot line to the secretory
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in protein folding. Annu. Rev. Biochem. 69:69-93.

o c

_ o

° o o o

° o

o o

° c

o o

Solutions

Chapter 2 Water

1. Hydrogen bonds involve strongly electronegative atoms such as nitrogen, oxygen, or sulfur.

(a)

— CHo — O

/ \

H H

(b)

0 H

(c)

H H

\ /

/ \

H H

— CH 2 — C — N
H

\ /

-CH-

N — H

/ \

H H

-O — H

2. (a) Glycerol is polar; it is not amphipathic; and it readily dissolves in water.

(b) Hexadecanoyl phosphate is polar; it is amphipathic; and it does not readily dissolve in
water but forms micelles.

(d) Glycine is polar; it is not amphipathic; and it readily dissolves in water.

3. There is a larger osmotic pressure inside the cells than outside because the molar concentra-
tion of solutes is much greater inside cells than outside. This results in a diffusion of water
into cells, causing them to swell and burst.

4 .

5 .

If the pH of a solution is below the piC a of any given ionizable group, the predominant species
will be the one with the dissociable proton on that group. If the pH of a solution is above the
piC a of any given ionizable group, the predominant species will be the one with the dissocia-
ble proton off of that group.

(a) pH = 11 where the — COO® form predominates.

(b) pH = 2 where the H® form predominates.

(d) pH =11 where the R — O® form predominates.

(a) Tomato juice. For pH = 4.2, if pH = —log [H®], then
[H®] = l(T pH [H®] = 10 -4 ' 2 = 6.3 X 1(T 5 M.

The ion-product constant of water (K w ) relates the concentrations of OH® and H®
(Equation 2.6).

[OH®] = KJ/[ H®] = 1.0 X 10~ 14 M 2 /6.3x -5 M = 1.6 X 10 -10 M.

(b) Human blood plasma. If the pH = 7.4, then

[H®] = 10 -7 - 4 = 4.0 X 10~ 8 M. [OH®] = iC w /[H®] =

1.0 X 10“ 14 M 2 /4.0x -8 M = 2.5 X 10 -7 M.

[H®] = 10 _1L6 = 2.5 X 10“ 12 M. [OH®] = K W /[H®] =
1.0 X 10~ 14 M 2 /2.0x -12 M = 4 X 10^ 3 M.

CH 2 — C' /vw '

697

698 SOLUTIONS Chapter 2

7. The total buffer species = [weak acid (HA)] + [conjugate base (A 0 )]

Total buffer concentration = 0.25 M + 0.15 M = 0.4 M

The pH can be calculated from the p K a and the concentrations given using the Henderson-
Hasselbalch equation.

[A 0 ] (0.15 M)

PH = p/C a + log— = 3.90 + log^^ = 3.90 - 0.22 = 3.68

8. The piC a for the ionization of H 2 PO 4 0 is 7.2. The Henderson-Hasselbalch equation (Equa-
tion 2.18) indicates that when the concentrations of the acidic form (H 2 PO 4 0 ) and its
conjugate base (HP0 4 ©) are equivalent, the pH is equal to the pl<f a , because the log term is
zero (log 1 = 0). Therefore, mixing 50 milliliters of solution A with 50 milliliters of solu-
tion B gives a buffer of pH 7.2. Since the concentration of each solution is 0.02 M, mixing
equal volumes gives a buffer whose phosphate concentration is also 0.02 M. The reason why
this is an effective buffer is that the final pH is at the p K a value. This means that the buffer will
resist changes in pH over a considerable range.

9. (a) The effective range of a buffer is from approximately one pH unit below to one pH unit

above the p fC a . The buffering range for MOPS is therefore 6. 2-8. 2, and the buffering
range for SHS is 4. 5-6. 5. Use the Henderson-Hasselbalch equation to calculate the ratios
of basic to acidic species.

For MOPS: pH

P K a + log

[RbN]

[R 3 NH©]

6.5 = 7.2 +

log

[RbN]

[R 3 NH©]

[RbN] 1

[R 3 NH©] 5

For SHS:

6.5

= 5.5 +

[RCOO 0 ]
109 [RCOOH]

[RCOO®] 10
[RCOOH] ~~ T

(b) An SHS buffer solution at pH 6.5 contains a much greater proportion of conjugate base
relative to acid (10:1) than MOPS does (1:5). Therefore, an SHS buffer would more effec-
tively maintain the pH upon addition of acid: H 0 + RCOO 0 < > RCOOH. Con-

versely, a MOPS buffer at pH 6.5 contains a greater proportion of acid than SHS does;
therefore, MOPS would more effectively maintain the pH upon addition of base:
R 3 NH© + OH 0 R 3 N + H 2 0.

10 .

pK a

1.2

■>

Partially ionized
(monoanion)

PKa

6.6

(dianion)

Second

endpoint

Chapter 2 SOLUTIONS

699

11 . Excess gaseous C0 2 rapidly equilibrates with aqueous C0 2 (Equation 2.25), leading to for-
mation of carbonic acid (Equation 2.23). Carbonic acid ionizes to H© and HC0 3 © (Equa-
tion 2.22). The excess acid, in the form of H©, can accumulate in bodily fluids, producing
acidosis.

12. Although the metabolism of lactate and other organic acids in the diet can lead to production
of C0 2 as shown, C0 2 is efficiently expired from the lungs (except during respiratory acido-
sis). Thus, the net product of the metabolic process is bicarbonate (HC0 3 ©), a base. Excess
H© present during metabolic acidosis can be removed when it combines with HC0 3 © to
form H 2 C0 3 (Equation 2.22), which then forms aqueous C0 2 and H 2 0 (Equation 2.23).

13. The acidic and conjugate base species of aspirin can be represented as RCOOH and RCOO©.
Use the Henderson-Hasselbalch equation to calculate the ratio of the two species at pH 2.0
and pH 5.0. Then calculate the fraction of the total species that is unionized and available for
absorption. In the stomach at pH 2.0,

2.0

P K a + log

[RCOO©]

[RCOOH]

3.5 +

log

[RCOO©]

[RCOOH]

The percentage of the uncharged species (RCOOH) is equal to the amount of RCOOH
divided by the total of RCOOH and RCOO©, times 100%.

[RCOOH]

X 1 00% =

1 + 0.03

X 100% = 97%

Therefore, nearly all aspirin in the stomach is in a form available for absorption. In the upper
intestine at pH 5.0, however, only a small percentage of aspirin is available for absorption.

[RCOO©]

5.0 = 3.5 + loq- -

y [RCOOH]

[RCOOH]

X 1 00% =

1 + 32

X 1 00% = 3%

Note that aspirin must be in solution in order to be absorbed. For this reason, coated or slow-
release forms of aspirin may alter the availability of aspirin in the stomach and intestine.

14. Use the Henderson-Hasselbach equation to calculate the ratio of the two species at each pH

At pH = 7.5

[H 2 NCH 2 CONH 2 ]

pH = p K a + log

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ]

7.5 = 8.2 + loq-

[ + H 3 NCH 2 CONH 2 ]

log

[H 2 NCH 2 CONH 2 ]

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ]

[ + H 3 NCH 2 CONH 2 ]

= 7.5 - 8.2 = -0.7

1 _

The ratio of [H 2 NCH 2 CONH 2 ] to [ + H 3 NCH 2 CONH 2 ] is 1 to 5. To determine the percent in
the conjugate base form: 1/(1 + 5)*100 = 17%. Therefore, 17% is unprotonated at pH 7.5.

At pH = 8.2
pH = p K a + log

[H 2 NCH 2 CONH 2 ]

[ + H 3 NCH 2 CONH 2 ]

700

SOLUTIONS Chapter 3

_ _ , , [H 2 NCH 2 CONH 2 ]

8.2 = 8.2 + loq-

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ]

log — = 8.2 - 8.2 = 0

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ] _ 1

[ + H 3 NCH 2 CONH 2 ] T

The ratio of [H 2 NCH 2 CONH 2 ] to [ + H 3 NCH 2 CONH 2 ] is 1.0 to 1.0. To determine the per-
cent in the conjugate base form: 1/(1 + 1)*100 = 50%. Therefore, 50% is unprotonated at
pH 8.2.

At pH 9.0:

, , [H 2 NCH 2 CONH 2 ]

pH = p K a + log—

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ]

9.0 = 8.2 + log-^ —

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ]

log-^ — = 9.0 - 8.2 = 0.8

[ + H 3 NCH 2 CONH 2 ]

[H 2 NCH 2 CONH 2 ] _ 6.3

[ + H 3 NCH 2 CONH 2 ] 1

The ratio of [H 2 NCH 2 CONH 2 ] to [ + H 3 NCH 2 CONH 2 ] is 6.3 to 1. To determine the percent
of the conjugate base: 63/(63 + 1)*100 = 86%. That is, 86% is unprotonated at pH 9.0.

15. This titration curve represents a compound with two p K a values, shown by the two plateaus
(near pH 2 and pH 10). Glycine has two p K a values at 2.4 and at 9.8.

16. Only (a) vitamin C would be soluble in water. Vitamin C contains several hydroxyl groups,
each of which can hydrogen-bond with water.

17. At 0°C the ion product for water is 1. 14 X 10~ 15 . At neutral pH,

At 100°C

Note that the density of water changes with temperature but this has very little effect
on [H + ].

18. HC1 dissociates completely in water. In 6 M HC1, [H®] = 6 M. The pH is — log(6) = —0.78.
The standard pH scale begins at zero ([H®] = 1 M) because it’s very unusual to encounter
more acidic solutions in biology.

Chapter 3 Amino Acids and the Primary Structures of Proteins

1. By comparing the priorities of L-cysteine (shown here) to those of L-serine (S configura-
tion, page 57) you will find that their sequence is clockwise and therefore L-cysteine has
the R configuration.

coo®

©h 3 n— c— h@

ch 2 sh

Chapter 3 SOLUTIONS 701

2. The stereochemistry of each chiral carbon must be examined to determine whether it has the
R or S configuration.

coo 0

©h 3 n— c—h@

CH(OH)CH 3

C-2, S-configuration

CH 3

C-3, ^-configuration

ch 2 ch 2 — nh 3 ci 0

, n © \

Cl 0 HN^NH

3. The other stereoisomers are:

coo 0

H — C — NH 3 0
HO — C— H

ch 3

D-Threonine

coo 0

NH 3 ® — C — H
HO — C — H

CH,

COO 0

H — C — NH 3 ®
HO— C — OH

CH,

L-Allothreonine D-Allothreonine

4. Methionine.

5. (a) Serine; phosphorylation of the hydroxyl group.

(b) Glutamate; carboxylation of the y-carbon.

6. By convention, peptides are designated from the N-terminus — > C-terminus, therefore Glu is
the N-terminus and Gly is the C-terminus.

COO v

i©

o ch 2

I II I

h 3 nch — ch 2 ch 2 c — NH — CH -

I y-Glu L

-NH-

-Cys-

-CH 2 -
- Gly -

-COO

7. The 6 residues at the C-terminus of melittin are highly hydrophilic (Table 3.1). Of the re-
maining 20 amino acid residues, nearly all are hydrophobic, including 9 with highly hy-
drophobic side chains (leucine, isoleucine, valine). The hydrophilic portion of melittin is
more soluble in aqueous solution, while the hydrophobic portion is more soluble in the
membrane lipids.

8. Use Table 3.2 to determine the net charge at each p K a value. The pH at which the net charge
is 0 lies midway between the two p K a values at which the average charges are +0.5 and —0.5.

(a) At pH 9.0, the net charge of arginine is +0.5, and at pH 12.5, the net charge is —0.5.
Therefore, pI Arg = (9.0 + 12.5) + 2 = 10.8.

(b) At pH 2.1, the net charge of glutamate is +0.5, and at pH 4.1, the net charge is —0.5.
Therefore, pI G i u = (2.1 + 4.1) -r- 2 = 3.1.

9. The ionizable groups are the free amino group of the N-terminal cysteine residue
(pJC a = 10.7), the glutamate side chain (p K a = 4.1), and the histidine side chain
(p K a = 6.0).

(a) At pH 2.0, the N-terminus and the histidine side chain have positive charges and the glu-
tamate side chain is uncharged. The net charge is +2.

(b) At pH 8.5, the N-terminus has a positive charge, the histidine side chain is uncharged,
and the glutamate side chain has a negative charge. The net charge is 0.

(c) At pH 10.7, the charge of the N-terminus is +0.5, the histidine side chain is uncharged,
and the glutamate side chain has a negative charge. The net charge is —0.5.

702 SOLUTIONS Chapter 3

10. (a)

ch 3 ch 3

x o/

s ch 2 o ch 3

(b) S

/ C '"N — H
N |

\ C /C-H

// I

o ch 2 oh

(c)

11. (a) Gly-Ala-Trp-Arg, Asp-Ala-Lys, Glu-Phe-Gly-Gln

(b) Gly-Ala-Trp, Arg-Asp-Ala-Lys-Glu-Phe, Gly-Gln

12. (a)

COOH

H 3 N — C — H

H®

pK a = 1.8

COO©

H 3 N — C — H

H®

P K a = 6.0

COO°

h 3 n — c — h

H®

pK a = 9.3

COO°

H 2 N — C — H

(b) A, 1; B, 3; C, 5; D, 7

(d) 4

(e) 5

(f) Histidine would be a good buffer within one pH unit of any of its three plC a values:
0.8-2. 8, 5. 0-7.0, and 8.3-10.3.

13. (a) Because there are two N- terminal groups, there must be two peptide chains, each having
an N-terminal aspartate residue.

(b) 2-Mercaptoethanol reduces disulfide bonds, and trypsin catalyzes cleavage on the car-
boxyl side of arginine residues. Since aspartate is found at both N-termini of FP, the se-
quence of the dipeptide is Asp-Arg, and the sequence of the pentapeptide is Asp-(Cys,
Gly, Met, Phe). The tripeptide has the sequence Cys-(Ala, Phe) and is derived from
trypsin-catalyzed cleavage of a pentapeptide whose sequence is Asp-Arg-Cys-(Ala, Phe).

(c) The C-terminal residue of each peptide chain is phenylalanine. Now that the terminal
residues are known, one peptide must have the sequence Asp-(Cys, Gly, Met)-Phe, and
the other must have the sequence Asp-Arg-Cys-Ala-Phe.

(d) CNBr cleaves on the carbonyl side of methionine residues to produce C-terminal ho-
moserine lactone residues. The peptides are therefore Asp-Met and (Cys, Gly)-Phe.
Glycine is the N-terminal residue of the tripeptide, so that pentapeptide sequence is
Asp-Met-Gly-Cys-Phe .

Chapter 3 SOLUTIONS

703

The complete FP structure is

Asp — Arg — Cys — Ala — Phe

Asp — Met — Gly — Cys — Phe

14. (a) The substitution of aspartate (D) for glutamate (E) at position 50 is an example of a con-

servative change. The amino acids aspartate and glutamate both contain acidic side
chains that are negatively charged at physiological pH.

(b) The substitution of tyrosine (Y) for histidine (H) is an example of a nonconservative
substitution since tyrosine contains an aromatic side chain and histidine contains a hy-
drophilic side chain consisting of an imidazole group.

15. The neurotransmitter serotonin is derived from the amino acid tryptophan.

In the conversion, the carboxyl group from tryptophan is removed and a hydroxyl group is
added to the aromatic ring.

Tryptophan

H,N® — C — H

Serotonin

16. (a) There are two peptide bonds present in TRH. They are marked with the dashed lines.

(b) TRH is derived from the tripeptide Glu-His-Pro. The proline carboxyl group has been
modified to an amide (marked with an*). The side chain carboxyl group of the amino
terminal Glu forms an amide with the residue’s cr-amino group (marked with a **).

o x ch 2
c ch 2

\ /

N — HC —
H

O O

II II

-c — NH — CH — c-

H?C

ch 2

/ \

CH,

\ / jr

-N — HC — C

NHo

Glu

= CH

His

17. (a) L-Dopa is in the S configuration.

(b) They are both derived from the amino acid tyrosine.

18. Although Figure 3.6 shows only three forms of alanine, there are actually four different forms
in equilibrium (see next page). The neutral form will be present at very low concentrations
because at any given pH the three other forms are much more stable. We can calculate the rel-
ative ratios of the four forms by assuming that the protonation/deprotonation of the two
charged groups is independent.

2.4 =

[R— COO 0 ]

2 ' 4 + '° 9 [R — COOH] theref ° re

[R— COO 0 ]
[R — COOH]

= 1

of H 3 N® — R to H 2 N — R is

= 9.9 + log [H * N ~ R] therefore [H * N ~ R] ~ 3.1 X 10 -
HH 3 f\P — R] [H 3 N 0 — R]

704

SOLUTIONS Chapter 3

ch 3

h 2 n— ch— coo°

H 3 N— CH— COOH

(neutral)

(cation)

Thus the relative ratios of the four forms are approximately

cation : zwitterion : anion : neutral 1:1: 10 -8 : 10 -8

and the concentration of the neutral form in a 0.01 M solution of alanine is about 10 -10 M.
Neutral molecules exist but their concentration is insignificant.

At pH 9.9 the ratios are

anion : zwitterion : cation : neutral 1:1: 10 -8 : 10 -8

At pH 6.15 the relative ratio of R — COO® and R — COOH is

[R — COO 0 ] [R — COO 0 ] ,

6.15 2.4 + log-- — — therefore — — 5.6 x 10 3

a [R — COOH] [R — COOH]

and the relative ratio of H 2 N — R and [H 3 N® — R] is

6.15

9.9 +

[H 2 N-R]
09 [H 3 N® — R]

[H 2 N — R]
[H 3 N® — R]

= 1.8 X 10“ 4

[H 3 N® — R]
[H 2 N — R]

= 5.6 X 1(T 8

The zwitterion is present in 5600-fold excess over the anion and cation forms and each of
these forms is 5600-fold more likely than the neutral form. The ratios are

zwitterion : anion : cation : neutral 3.1 x 10 7 : 1 : 1.8 x 1(T 4 : 1.8 x 1(T 4 : 3.2 X 1CT 8

The concentration of the neutral form in a solution of 0.01 M alanine is insignificant.

19. The relative concentrations of the zwitterion and the cation are

2.4 = 2.4 + log

[H 3 N-

ch 3

-c — COO°]

[H 3 N-

ch 3

-c — COO°]

ch 3

[H 3 N— C-COOH] [H 3 N-C — COOH]

= 1

Thus the concentration of the zwitterion in a solution of 0.01 M alanine is 0.005 M. (We can
ignore the concentrations of the anion and neutral forms — see previous question.)

At pH 4.0

4.0

2.4 +

log

[zwitterion]

[cation]

The concentration of the zwitterion is 0.01 M X f?

[zwitterion]

[cation]

0.00976 M

Chapter 4 SOLUTIONS

705

Chapter 4 Proteins: Three-Dimensional Structure and Function

(b) The R groups represent the side chains of the amino acid residues.

(d) Both peptide groups in this tripeptide are in the trans conformation, since the a-carbon
atoms are on opposite sides of the peptide bonds.

(e) The peptide groups may rotate around the N — C a and C a — C bonds.

2. (a) (1) In an a helix, intrachain hydrogen bonds form between carbonyl oxygens of certain

residues and amide hydrogens of other residues. The hydrogen bonds are approxi-
mately parallel to the helix axis (Figure 4.10).

(2) In a collagen triple helix, interchain hydrogen bonds form between amide hydrogens
of the glycines in one chain and carbonyl oxygen atoms of residues (which are often
proline) in an adjacent chain (Figure 4.41). There are no intrachain hydrogen bonds
in a collagen helix.

(b) The side chains of an a helix point outward from the cylinder of the helix (Figure 4.11).
In collagen, three chains coil around each other so that every third residue of a given
chain makes contact with the other two chains along the central axis of the triple helix
(Figure 4.42). Only the small side chain of glycine can fit at these positions. The other
side chains point outward from the triple helical coil.

3. (1) The presence of glycine in an a helix destabilizes the helix due to the greater freedom of

movement allowed by the small side chain. For this reason, many a helices begin or end
with glycine.

(2) Proline tends to disrupt a helices because its rigid, cyclic side chain stereochemically in-
terferes with the space that would normally be occupied by a neighboring residue in the
a helix. In addition, proline lacks a hydrogen on its amide nitrogen and cannot participate
in normal intrahelical hydrogen bonding.

4. (a) Due to the flexibility resulting from a small side chain ( — H), glycine is often found in

“hairpin loops” that connect sequential antiparallel /3 strands. The glycine residues (G) in
positions 8 and 14 provide two hairpin-loop regions to connect the three f3 strands in Be-
tanova.

®— 1

r<^

C [ ^COOQ

(b) [3 -sheet structures are stabilized by hydrogen bonds that form between a carbonyl oxy-
gen of one strand and an amide nitrogen of an adjacent strand (Figure 4.15).

h 3 n— c —

0
H

COO 0

5. Helix-loop -helix (HLH) motif (Figure 4.19).

6. (a) a//3. Regions of a helix and /3 strand alternate in the polypeptide chain.

(b) al (3 barrel. Parallel /3 strands are surrounded by a layer of a helices in a cylindrical
shape.

(c) Yeast FMN oxidoreductase and E. coli enzyme required for tryptophan biosynthesis
(Figure 4.24 (i) and (j) respectively).

706

SOLUTIONS Chapter 4

7. Protein disulfide isomerase contains two reduced cysteine residues at the active site, and these
participate in a reduction and disulfide exchange that allows the misfolded protein to refold
into the lower energy native conformation.

8. The highly hydrophobic side chains of methionine, leucine, phenylalanine, and isoleucine are
most likely to be on the side of the helix that faces the interior of the protein. Most of the
other side chains are polar or charged and can interact with the aqueous solvent. Since the
a helix is a repeating structure with approximately 3.6 residues per turn, the hydrophobic
groups must be found every three or four residues along the sequence, so that one side of the
helix is hydrophobic.

9. Covalent cross-linking contributes significantly to the strength and rigidity of collagen fibers.
In one type of cross-link, allysine residues in a collagen molecule condense with lysine
residues in an adjacent molecule, forming Schiff bases (Figure 4.38a). When an allysine
residue reacts with homocysteine, it is unable to participate in the normal cross-linking of
collagen molecules. High levels of homocysteine in blood probably lead to defective collagen
structure and skeletal deformities.

0 = C

i //

CH — CH 2 — CH 2 — CH 2 — C

I \

HN H

s Allysine residue

* in collagen

coo©

H 2 N — CH — CH 2 — CH 2 — SH
Homocysteine

= c coo©

I I

CH — CH 2 — CH 2 — CH 2 — CH = N — CH — CH 2 — CH 2 — SH
HN

10. The sequence -Gly-Pro-X-Y- occurs frequently in collagen, which is found throughout the
body, including in the skin. Because the larval enzyme can catalyze cleavage of collagen
chains, the parasite is able to enter the host.

11. The reaction of carbon dioxide with water explains why there is a concomitant lowering of
pH when the concentration of C0 2 increases. Carbon dioxide produced by rapidly metabo-
lizing tissue reacts with water to produce bicarbonate ions and H® .

The H© generated in this reaction decreases the pH of the blood and thus stabilizes the
deoxy form (T conformation) of hemoglobin. The net effect is an increase in the P 50 > that is,
a lower affinity of hemoglobin for oxygen, so that more oxygen is released to the tissue
(Figure 4.50). Carbon dioxide also lowers the affinity of hemoglobin for oxygen by forming

Chapter 5 SOLUTIONS

707

carbamate adducts with the N-termini of the four chains (Figure 4.51). These adducts
contribute to the stability of the deoxy (T) conformation, thereby further increasing the
P 50 and promoting the release of oxygen to the tissue.

(b) Shock victims suffer a critical deficit of oxygen supply to their tissues. Bicarbonate ad-
ministered intravenously provides a source of carbon dioxide to the tissues. By lowering
the affinity of hemoglobin for oxygen, carbon dioxide facilitates a release of oxygen from
oxyhemoglobin to the tissues.

12. (a) 2,3BPG binds to positively charged side chains in the central cavity of deoxyhemoglobin

(Figure 4.49). Since Hb F lacks two positively charged groups (His- 143 of each (3 chain),
2,3BPG binds less tightly to Hb F than to Hb A.

(b) 2,3BPG stabilizes the deoxy form of hemoglobin, increasing the fraction of molecules in
the deoxy form. Since Hb F binds 2,3BPG less tightly than does Hb A, Hb F is less af-
fected by 2,3BPG in the blood and has a greater fraction of molecules in the oxy form. Hb
F therefore has a greater affinity than Hb A for oxygen at any oxygen pressure.

(c) At the oxygen pressure of tissues, 20-40 torr, Hb F has a greater affinity for oxygen than
does Hb A. The difference in affinity allows efficient transfer of oxygen from maternal
blood to the fetus.

13. The low P 50 value of Hbyakima indicates a greater than normal affinity for oxygen even at the
oxygen pressures found in working muscle. The increased affinity means that Hbyakima gives
up less oxygen to the working muscle.

14. (a) Hydrophilic (italicized) and hydrophobic (underlined) residues are identified:

ECG KFMWK CKNSNDCCKDYV CSSRWKW CVLA5PF
(b) In the three-dimensional structure of proteins, amino acids that are far from each other
in the primary sequence can interact in the globular structure of the protein. Thus the
hydrophobic amino acids can be very close to each other in the three-dimensional struc-
ture and provide a “hydrophobic” face for interaction with the membrane.

15. (a) The most effective binding of selenoprotein P to heparin is seen at a pH below 6. The

binding of selenoprotein P to heparin decreases as the pH is increased to 7. There is very
little binding of selenoprotein P to heparin at pH values greater than 7.

(b) Heparin is negatively charged. If selenoprotein P is positively charged, it can bind to he-
parin. Histidine residues are abundant in selenoprotein P. Histidine has an imidazole side
chain that has a p K a value of 6.0. That is, at a pH of 6.0, 50% of the histidine residues
would be protonated and positively charged and 50% would be unprotonated and un-
charged. Below a pH of 6.0, there would be a net positive charge on the histidine residues,
resulting in effective electrostatic interactions with the heparin. At pH values above 7, al-
most all of the histidine residues would be unprotonated and uncharged and will not ef-
fectively interact with the negatively charged heparin molecule.

16. Collagen is protein consisting of three polypeptide chains that are wound together in a triple
helix. The protease bromelin is an enzyme that cleaves some of the peptide bonds in the
polypeptide chains. The polypeptide chains are necessary to trap the water molecules in a
semisolid state when gelatin cools, and if these are cleaved, the gelatin will not set properly.
The cleavage of the polypeptide chains in collagen by bromelin destroys the ability of the gel-
atin to harden. If the pineapple is first cooked, the heat will denature the protein and thus the
enzyme activity will be destroyed. Therefore cooked pineapple can be added to slightly thick-
ened gelatin, and the gelatin will proceed to the semisolid state as desired. (Assume that heat
denaturation is irreversible.)

17. The replacement of lysine by methionine results in one less positive charge on each beta sub-
unit in the central cavity (see Figure 4.49). 2,3BPG binds less tightly to HbH. This causes
more of the mutant protein to be in the R state (oxyhemoglobin is stabilized). The curve is
shifted towards the left (more like myoglobin). Since more is in the R state, the affinity for
oxygen has increased.

Chapter 5 Properties of Enzymes

1. The initial velocities are approaching a constant value at the higher substrate concentrations,
so we can estimate the V max as 70 mM/min. Since K m equals the concentration of substrate
[S] required to reach half the maximum velocity, we can estimate the K m to be 0.01 M since
that’s the concentration of substrate that yields a rate of 35 mM/min (= ^^>2).

2. (a) The ratio k cat /K m , or specificity constant , is a measure of the preference of an enzyme for

different substrates. When two substrates at the same concentration compete for the

708 SOLUTIONS Chapter 5

active site of an enzyme, the ratio of their rates of conversion to product is equal to the
ratio of the k cat /K m values, since v 0 = (k cat /K m ) [E] [S] for each substrate and [E] and [S]
are the same.

V-o(Sl) (^ca t >^m) 1 >[E][S]

v o(S 2 ) (/c cat >/C m ) 2 >[E][S]

(b) The upper limit of k cat /K m approaches 10 8 to 10 9 s -1 , the fastest rate at which two un-
charged molecules can approach each other by diffusion at physiological temperatures.

(c) The catalytic efficiency of an enzyme cannot exceed the rate for the formation of ES from
E and S. The most efficient enzymes have k cat /K m values approaching the rate at which
they encounter a substrate. At this limiting velocity they have become as efficient catalysts
as possible because every encounter produces a reaction. (Most enzymes don’t need to
catalyze reactions at the maximum possible rates so there’s no selective pressure to evolve
catalytically perfect enzymes.)

3. The catalytic constant (fc cat ) is the first-order rate constant for the conversion of ES to
E + P under saturating substrate concentrations (Equation 5.26), and CA has a much
higher catalytic activity in converting substrate to product than does OMPD. However, the
efficiency of an enzyme can also be measured by the rate acceleration provided by the en-
zyme over the corresponding uncatalyzed reaction (/c cat //c n , Table 5.2). The reaction of the
substrate for OMPD in the absence of enzyme is very slow (k n = 3 X 1CT 16 s -1 ) compared
to the reaction for the CA substrate in the absence of enzyme {k n = IX 10 -1 s -1 ). There-
fore, while the OMPD reaction is much slower than the CA reaction in terms of /c cat ,
OMPD is one of the most efficient enzymes known and provides a much higher rate accel-
eration than does CA when the reactions of each enzyme are compared to the correspon-
ding uncatalyzed reactions.

4. When [S] = 100 /xM, [S] » K mi so v 0 = V max = 0.1 /ulM min -1 .

(a) For any substrate concentration greater than 100 /xM, v 0 = V max = 0.1 /xM min -1 .

(b) When [S] = K m , v 0 = V m2 Jl, or 0.05 /ulM min -1 .

(c) Since K m and V^x are known, the Michaelis-Menten equation can be used to calculate
v 0 at any substrate concentration. For [S] = 2 /xM,

Knax[S]

/C m + [S]

(0.1 ^iM min -1 )(2 ^M)
(1 + 2 yuM)

0.2

/jlW\ min

0.067 min 1

5. (a) Determine [E] tota j in moles per liter, then calculate V max .

- 0.2 g - 9.3 X ,° * M

Knax = *cat[E]total = 1000 S" 1 ^ X 10“ 6 M) = 9.3 X 1 0“ 3 M S^ 1

(b) Since V max is unchanged in the presence of the inhibitor, competitive inhibition is
occurring. Because the inhibitor closely resembles the heptapeptide substrate, compet-
itive inhibition by binding to the enzyme active site is expected (i.e., classical competi-
tive inhibition).

6. Curve A represents the reaction in the absence of inhibitors. In the presence of a competitive
inhibitor (curve B), K m increases and V max is unchanged. In the presence of a noncompetitive
inhibitor (curve C), V max decreases and K m is unchanged.

[S]

Chapter 5 SOLUTIONS

709

7. Since the inhibitor sulfonamides structurally resemble the PABA substrate we would predict
that sulfonamides bind to the enzyme active site in place of PABA and act as competitive
inhibitors (Figure 5.9).

8. (a) To plot the kinetic data for fumarase, first calculate the reciprocals of substrate concentra-
tions and initial rates of product formation. (Note the importance of including correct
units in calculating and plotting the data.)

Fumarate ^ Rate of product formation

[S] (mM) — (mM -1 ) v 0 (mmol I -1 min -1 ) —(mmol -1 I min)

2.0

0.50

2.5

0.40

3.3

0.30

3.1

0.32

5.0

0.20

3.6

0.28

10.0

0.10

4.2

0.24

Vmax i s obtained by taking the reciprocal of l/V max from the y intercept (Figure 5.6).

1/V max = 0.20 mmol -1 I min, so V max = 5.0 mmol I -1 min -1

K m is obtained by taking the reciprocal of — VK m from the v intercept.

-VK m = -0.5 mivr 1 , so K m = 2.0 mM or 2 X 10~ 3 M

(b) The value of k cat represents the number of reactions per second that one enzyme active
site can catalyze. Although the concentration of enzyme is 1 X 10 -8 M, fumarase is a
tetramer with four active sites per molecule so the total concentration of enzyme active
sites [E tota i] is 4 X 10 -8 M. Using Equation 5.26:

Vmax _ 5.0 mmol I 1 min 1 1 min _

[Etotai] 4 X 10 -5 mmol I -1 60 s

9. Like pyruvate dehydrogenase (PDH) (Figure 5.22), glycogen phosphorylase (GP) activity is
regulated by alternate phosphorylation by a kinase and dephosphorylation by a phosphatase.
However, unlike PDH, the active form of GP has two phosphorylated serine residues; in the
inactive GP form, two serine residues are not phosphorylated.

710

SOLUTIONS Chapter 5

10. Inhibition of the first committed step of a multistep pathway allows the pathway to proceed
only when the end product is needed. Since the first committed step is regulated, flux in the
pathway is controlled. This type of regulation conserves raw material and energy.

11. When [aspartate] = 5mM,v 0 = V max /2. Therefore, in the absence of allosteric modulators,
K m = [S] = 5 mM. ATP increases v 0 , and CTP decreases v 0 .

12. (a) To plot the kinetic data for P450 3A4, first calculate the reciprocals of substrate concen-
trations and initial rates of product formation. The data are plotted in the double recip-
rocal plot and are shown with the dashed line.

Midazolam

ISK/iM)

i/rs] ( m m-')

Rate of product formation
v 0 (pmol I -1 min -1 )

1/y 0 (pmol 1 1 min)

100

0.01

0.5

156

0.0064

0.25

222

0.0045

0.125

323

0.0031

Vmax i s obtained by taking the reciprocal of 1/V max from the y intercept (Figure 5.6).
1/Knax ~~ 0.0025 pmol -1 I min, so V max = 400 pmol I -1 min^ 1

Chapter 6 SOLUTIONS

711

K m is obtained by taking the reciprocal of — 1A K m from the x intercept

-1 /K m = -0.3 /tM -1 , so K m = 3.3

(b) The reciprocals of the substrate concentration and activity in the presence of ketocona-
zole are given in the table.

Midazolam
IS] (pM)

1/IS] (jliM- 1 )

Rate of product formation
in the presence of
0.1 /jlM ketoconazole/
y 0 (pmol I -1 min -1 )

1/iz 0 (pmol 1 1 min)

0.091

0.5

0.056

0.25

0.037

0.125

0.025

The plot of the data (solid line) is given in the double reciprocal plot shown in (a). There is
an increase in the y intercept and no apparent change in the x intercept. From the double re-
ciprocal plot, it appears that ketoconazole is a noncompetitive inhibitor (see Figure 5.11).
These inhibitors are characterized by an apparent decrease in V max (increase in 1/V max ) with
no change in K m .

13 . (a) Bergamottin appears to inhibit the activity of P450 3A4 since the P450 activity measured

in the presence of 0.1 and 5 fiM bergamottin is less than that of the P450 activity in the
absence of bergamottin.

(b) It might be dangerous for a patient to take their medication with grapefruit juice since
there appears to be an inhibition of P450 activity in the presence of bergamottin. If the
bergamottin decreases the P450 activity, and the P450 enzyme is known to metabolize
the drug to an inactive form, the time it takes to convert the drug to its inactive form may
be increased. This may prolong the effects of the drug, which may lead to adverse conse-
quences for the patient.

14 . (a) When [S] » K m , then K m + [S] ~ [S]. Substrate concentration has no effect on velocity,

and v 0 = V^x, as shown in the upper part of the curve in Figure 5.4a.

= WnaxtS] WS]

V ° K m + [S] ~ [S]

(b) When [S] K m , K m + [S] ~ K m , and the Michaelis-Menten equation simplifies to

W s] , \WS]

V ° K m + [S] ~ K m

Velocity is related to [S] by a constant value, and the reaction is first order with respect to
S, as shown in the lower part of the curve in Figure 5.4a.

^max _ Knax[S]

V ° “ 2 ~ K m + [S]

K m + [S] = 2[S]

Km = [S]

Chapter 6 Mechanisms of Enzymes

1 . (a) The major binding forces in ES complexes include charge-charge interactions, hydrogen
bonds, hydrophobic interactions, and van der Waals forces. (About 20% of enzymes bind
a substrate molecule or part of it covalently.)

(b) Tight binding of a substrate would produce an ES complex that lies in a thermodynamic
pit, effectively increasing the activation energy and thereby slowing down the reaction.
Tight binding of the transition state, however, lowers the energy of the ES^ complex,
thereby decreasing the activation energy and increasing the rate of the reaction.

712

SOLUTIONS Chapter 6

2. The activation barrier for the reaction is lowered by (1) raising the ground-state energy level
(ES) and (2) lowering the transition-state energy level (ES*), resulting in a reaction rate in-
crease.

3. The rate determining step of a multistep reaction is the slowest step, which is the step with the
highest activation energy. For Reaction 1, Step 2 is the rate determining step. For Reaction 2,
Step 1 is the rate determining step.

4. The reactive groups in Reaction 2 ( — OH and — COOH) are held at close proximity. They
are oriented in a manner suitable for catalysis by steric crowding of the bulky methyl groups
of the ring. The reactive — COOH group cannot rotate away as freely as it can in Reaction 1.
Model systems such as these are relevant because they indicate potential rate increases that
might be obtained by enzymes that bring substrates and the enzymes catalytic groups into
positions that are optimal for reaction.

5. (1) Binding effects. Fysozyme binds the substrate so that the glycosidic bond to be cleaved is

very close to both of the enzyme catalytic groups (Glu-35 and Asp-52). In addition, the
energy of the ground- state sugar ring is raised because it is distorted into a half- chair con-
formation.

(2) Acid-base catalysis. Glu-35 first donates a proton to an oxygen of the leaving sugar (gen-
eral acid catalysis), and then accepts a proton from the attacking water molecule (general
base catalysis).

(3) Transition- state stabilization. Asp-52 stabilizes the developing positive charge on the ox-
ocarbocation intermediate, and subsite D favors the half-chair sugar conformation of
this intermediate. The structure proposed for the transition state includes both this
charge and sugar conformation in addition to hydrogen bonding to several active- site
residues.

6. Serine 195 is the only serine residue in the enzyme that participates in the catalytic triad at the
active site of cr-chymotrypsin. The resulting increase in the nucleophilic character of Ser-195
oxygen allows it to react rapidly with DFR

7. (a) The catalytic triad is composed of an aspartate, a histidine, and a serine residue. Histidine

acts as a general acid-base catalyst, removing a proton from serine to make serine a more
powerful nucleophile in the initial step. Aspartate forms a low-barrier hydrogen bond
with histidine, stabilizing the transition state. An acid catalyst, histidine donates a proton
to generate the leaving amine group.

(b) The oxyanion hole contains backbone — NH — groups that form hydrogen bonds with
the negatively charged oxygen of the tetrahedral intermediate. The oxyanion hole medi-
ates transition-state stabilization since it binds the transition state more tightly than it
binds the substrate.

(c) During catalysis, aspartate forms a low-barrier hydrogen bond with the imidazolium
form of histidine. Because asparagine lacks a carboxylate group to form the stabilizing
hydrogen bond with histidine, enzyme activity is dramatically decreased.

Chapter 6 SOLUTIONS

713

8. (a) Human cytomegalovirus protease: His, His, Ser (b) /3- Lactamase: Glu, Lys, Ser

Glu Lys Ser

I n I 1

H 2

Asp Lys Thr

1 n 1 1

H 3

9. When tyrosine was mutated to phenylalanine, the activity of the mutant enzyme was less than
1% of the wild-type enzyme. Thus, the tyrosine residue is involved in the catalytic activity of
DDP-IV. Tyrosine contains an -OH group on the aromatic ring of the side chain. As previously
stated, this tyrosine is found in the oxyanion hole of the active site. Hydrogen bonds in the
oxyanion hole of serine proteases are known to stabilize the tetrahedral intermediate. Tyro-
sine with an -OH group on the side chain can form a hydrogen bond and stabilize the tetra-
hedral intermediate. Phenylalanine does not have a side chain that can form a hydrogen
bond. Therefore, the tetrahedral intermediate will not be stabilized resulting in a loss of
enzyme activity.

10. (a) Acetylcholinesterase catalytic triad: Glu-His-Ser

His

Glu \ Ser

1 n H i

(d) Hepatitis A protease:

His

Asp

1 n 1

COO 0 H — O HN /N:-

His

Ser

HN .N HN .N: HO — CH 2

(b)

Ser

-CH 2

\*e

i-PrO OO-U

H©

0°© o-

>n\-v

f—

Ser

-ch 2

i-PrO

OCH,

Ser

°\/°

i-PrO OCH

ch 2

11. Transition-state analogs bound to carrier proteins are used as antigens to induce the formation
of antibodies with catalytic activity. The tetrahedral phosphonate ester molecule is an analog
of the tetrahedral intermediate structure in the transition state for hydrolysis of the benzyl
ester moiety of cocaine. An antibody raised against the phosphonate structure that was able to
stabilize the transition state of the cocaine benzyl ester hydrolysis could effectively catalyze this
reaction.

12. (a) Wild-type a \ -proteinase inhibitor is given as treatment to individuals who produce an
a 1 -proteinase inhibitor with substitutions in the amino acid sequence. These changes re-
sult in a protein that does not effectively inhibit the protease elastase. Uncontrolled elas-
tase activity leads to increased breakdown of elastin, leading to destructive lung disease.
Therefore, these patients are given a functional elastase inhibitor.

Cys

-HS — CH 2

+ F 0

714

SOLUTIONS Chapter 7

(b) The treatment for al -proteinase inhibitor deficiency is to administer the wild-type pro-
tein intravenously. If the protein is given orally, the enzymes present in the digestive tract
will cleave the peptide bonds in the a 1 -proteinase inhibitor. By administering the drug
directly into the bloodstream, the protein can circulate to the lungs to act at the site of the
neutrophil elastase.

Chapter 7 Coenzymes and Vitamins

1. (a) Oxidation; NAD®, FAD, or FMN. (The coenzyme for the reaction shown is NAD®.)

(b) Decarboxylation of an a-keto acid; thiamine pyrophosphate.

(d) Molecular rearrangement; adenosylcobalamin.

(e) Transfer of a hydroxyethyl group from TDP to CoA as an acyl group; lipoic acid.

2. (a) NAD®, NADP®, FAD, FMN, lipoamide, ubiquinone. Protein coenzymes such as thiore-

doxin and the cytochromes.

(b) Coenzyme A, lipoamide.

(d) Pyridoxal phosphate

(e) Biotin, thiamine pyrophosphate, vitamin K

3. No. NAD® acquires two electrons but only one proton. The second proton is released into
solution and is reutilized by other proton-requiring reactions.

5. NAD®, FAD, and coenzyme A all contain an ADP group (or ADP with 3 '-phosphate for
coenzyme A).

7. Vitamin B 6 is converted to pyridoxal phosphate, which is the coenzyme for a large number of
reactions involving amino acids, including the decarboxylation reactions in the pathways that
produce serotonin and norepinephrine from tryptophan and tyrosine, respectively. Insuffi-
cient vitamin B 6 can lead to decreased levels of PLP and a decrease in the synthesis of the neu-
rotransmitters.

8. The synthesis of thymidylate (dTMP) requires a tetrahydrofolate (folic acid) derivative. Defi-
ciency of folic acid decreases the amount of dTMP available for the synthesis of DNA. De-
creased DNA synthesis in red blood cell precursors results in slower cell division, producing
macrocytic red blood cells. The loss of cells by rupturing causes anemia.

9. (a) Cobalamin.

(b) The cobalamin derivative adenosylcobalamin is a coenzyme for the intramolecular re-
arrangement of methylmalonyl CoA to succinyl CoA (Figure 7.28). A deficiency of
adenosylcobalamin results in increased levels of methylmalonyl CoA and its hydrolysis
product, methylmalonic acid. Another cobalamin derivative, methylcobalamin, is a coen-

Chapter 7 SOLUTIONS 715

zyme for the synthesis of methionine from homocysteine (Reaction 7.5), and a deficiency
of cobalamin results in an excess of homocysteine and a deficiency of methionine.

10 . (a) In one proposed mechanism, a water molecule bound to the zinc ion of alcohol dehydro-
genase forms OH®, in the same manner as the water bound to carbonic anhydrase
(Figure 7.2). The basic hydroxide ion abstracts the proton from the hydroxyl group of
ethanol to form H 2 0. (Another mechanism proposes that the zinc also binds to the alco-
holic oxygen of the ethanol, polarizing it.)

(b) No, a residue such as arginine is not required. Ethanol, unlike lactate, lacks a carboxylate
group that can bind electrostatically to the arginine side chain.

11. A carboxyl group is transferred from methylmalonyl CoA to biotin to form carboxybiotin
and propionyl CoA.

12. (a)

Carboxybiotin

HN^N

W H

CH 2 ^ | ^COO 0

(b) Racemization would not occur. Although a Schiff base forms during decarboxylation as
well as racemization, the reactive groups in the histidine decarboxylase active site specifi-
cally catalyze decarboxylation, not racemization, of histidine.

716 SOLUTIONS Chapter 8

13. (a) See Reactions 13.2-13.4 on pages 412 and 413.

(c)

(b)

CHo

HETDP

TDP

— CH-
+

Acetyl-TDP

TDP

-OH

CH,

-C = 0

TPP

H ,c-c-s

Lipoamide

Dihydrolipoamide Acetyl-dihydrolipoamide

TDP

hoch 2 — c— oh

HC = 0

H — C— OH

Chapter 8 Carbohydrates

1. (a) D-Glucose and D-mannose

(b) L-Galactose

(d) Dihydroxyacetone

2. (a)

^TDP

' A

- ' ru

HOCH 2 — C-^OH
HOC — H

H — C — OH

H — C— OH

CHyOPOP

CH 2 OH
C = 0

HOCH

H — C — OH

H — C— OH

CH20P03®

TPP

(e) Erythrulose (either D or l)

(f) D-Glucose

(g) N-Acetylglucosamine

H .0

(b) H O

\ < J r

(c)

H —

CH 2 OH
C — OH

< d > \/>
c
1

H — C— OH
|

HO— C — H
|

HO —

C — H

HO— C — H

H — C— OH

H — C — OH

H —

C — OH

H — C— OH

HO— C — H

-u—

ch 2 oh

HO— C — H
|

-u-

H — C — OH

CH 2 OH

ch 3

1 ©

COO^

3. Glycosaminoglycans are unbranched heteroglycans of repeating disaccharide units. One
component of the disaccharide is an amino sugar and the other component is usually an al-
duronic acid. Specific hydroxyl and amino groups of many glycosaminoglycans are sulfated

(a)

CH 2 OH

/3-D-Fructofuranose

5. (a) a-Anomer

(b) Yes, it will mutorotate.

(b)

CH 2 OH

/3-D-Fructopyranose

(d) A ketone

(e) Four chiral carbons

Chapter 8 SOLUTIONS 717

6. Glucopyranose has five chiral carbons and 2 5 , or 32, possible stereoisomers; 16 are D sugars
and 16 are L sugars. Fructofuranose has four chiral carbons and 2 4 , or 16, possible stereoiso-
mers; 8 are D sugars and 8 are L sugars.

( d ) CHO°

HO — C — H

H — C — OH

HO — C — H

COO

8. Only the open-chain forms of aldoses have free aldehyde groups that can form Schiff bases
with amino groups of proteins. Because relatively few molecules of D-glucose are found in the
open-chain form, D-glucose is less likely than other aldoses to react with proteins.

9. A pyranose is most stable when the bulkiest ring substituents are equatorial, minimizing
steric repulsion. In the most stable conformer of /3-D-glucopyranose, all the hydroxyl groups
and the — CH 2 OH group are equatorial; in the most stable conformer of cr-D-glucopyranose,
the C- 1 hydroxyl group is axial.

10. O

Envelope

conformation

11. The a and (3 anomers of glucose are in rapid equilibrium. As (3- D-glucose is depleted by the
glucose oxidase reaction, more (3 anomer is formed from the a anomer until all the glucose
has been converted to gluconolactone.

12. Sucralose is a derivative of the disaccharide sucrose (see Figure 8.20). The two hydroxyl
groups on C-l and C-6 of the fructose molecule have been replaced with chlorine. The hy-
droxyl group on C-4 of the glucose molecule was removed and then chlorine added. In the
chemical synthesis of sucralose from sugar, the configuration of the C-4 substituent of the
glucose moiety is reversed.

718 SOLUTIONS Chapter 9

15. (a) a, b, and c; these oligosaccharides contain GlcNAc — Asn bonds.

(b) b and c; these oligosaccharides contain /3-galactosidic bonds.

(d) None, since none of the oligosaccharides shown contains fucose.

, i i , i

a(2->3) [3 (1— >4)

linkage linkage

17. Paper is made of cellulose and /3-glucosidases break down cellulose to glucose residues. If you
took a pill, this book would still taste like chewed up paper that tastes like paste (ugh!). That’s
because your taste buds are in your mouth and the enzyme is in your stomach. If you mari-
nate the book in an enzyme solution, it would taste much sweeter.

Publishers would not print textbooks using flavored ink because they, and the authors, want
students to keep their textbooks as valuable resources for future reference in the many ad-
vanced courses that you are planning to take. On the other hand, encouraging students to eat
their textbooks, instead of selling them, might be a good thing because it promotes better
health and nutrition.

(a)

H H

I I

CH 3 (CH 2 ) 7 — c = c
( a) CH 3 CH 2 CH 2

Chapter 9

2 .

Lipids and Membranes

(b) H H

| |

(c)

ch 2

/ V

3. (a) co- 3; (b) co-6; (c) co-6 ; (d) neither (co- 9); (e) co-6.

4 .

HoC— CH— CH

cn 3

©I

2 n —

CH,

Chapter 9 SOLUTIONS 719

(a)

(b) Docosahexaenoic acid is classified as an co-3 fatty acid

6 .

_ II /

(R,) (R 2 )

nh 3

I \

coo©

(R,)

A lysolecithin

Fatty acid

7. (a)

1 2 3

nh 3

ch 2

1 ©
0 = P— o u

'I I

? ?

0 = C C = 0

I I

(h 2 c) 16 (ch 2 ) 7

I I

CH, C — H

C — H

(b) CH 3
©I

H 3 C — N — CH 3

ch 2

0 = P — o

C — H

0=C

(H 2 C),.

H — C

(CH 2 ) l:

CH 3 CH 3

(ch 2 ) 7

ch 3

720

SOLUTIONS Chapter 9

8. (a)

ch 2 oh

c=o

13 .

14 .

15 .

16 .

9 . PE contains docosahexaenoic acid at position C-2 on the glycerol- 3 -phosphate backbone at
both temperatures. At lower temperatures, the percent of the monounsaturated fatty acyl
groups at position C-l increased from 14% at 30°C to 39% at 10°C. The membrane fluidity
must be maintained for the organism, and this is accomplished by changing the composition
of the membrane lipids. The increase in the unsaturated lipids at the lower temperature will
allow for the proper membrane fluidity.

10. Farnesyl transferase adds a farnesyl or “prenyl” group to a cysteine side chain of the ras pro-
tein (Figure 9.23b). The ras protein is subsequently anchored to the plasma and endoplasmic
reticulum membranes and is active in cell signaling processes. Farnesyl transferase is a
chemotherapy target because inhibition of this enzyme in tumor cells would disrupt the sig-
naling activity of the mutated ras protein. In fact, farnesyl transferase (FT) inhibitors are po-
tent suppressors of tumor growth in mice.

11. Fine A represents diffusion of glucose through a channel or pore, and line B represents pas-
sive transport. Diffusion through a channel or pore is generally not saturable, with the rate
increasing linearly with the concentration of the solute. Transport via a transport protein is
saturable at high solute concentrations, much like an enzyme is saturated at high substrate
concentrations (Section 9. 10C).

12 .

HCI

Theobromine is structurally related to caffeine and theophylline (Figure 9.45). The methylated
purines, including the obromine, inhibit cAMP phophodiesterase, a soluble enzyme that cat-
alyzes the hydrolysis of cAMP to AMP (Figure 9.43). These methylated purines inhibit the
breakdown of the intracellular messenger cAMP to AMP. Therefore, the effects of the cAMP
are prolonged. For dogs, this is combined with the fact that they have slower clearance of the
ingested theobromine from their system. Both of these result in the toxicity associated with
ingesting the chocolate.

The two second messengers IP 3 and DAG are complementary in that they both promote the
activation of cellular kinases, which then activate intracellular target proteins by causing their
phosphorylation. Diacylglycerol activates protein kinase C directly, whereas IP 3 elevates Ca©
levels by opening a Ca© channel in the membrane of the endoplasmic reticulum, releasing
stored Ca© into the cytosol (Figure 9.48). The increased Ca© levels activate other kinase
leading to a phosphorylation and activation of certain target proteins.

Insulin can still bind normally to the a subunits of the insulin receptor, but due to the muta-
tion, the (3 subunits lack tyrosine-kinase activity and cannot catalyze autophosphorylation or
other phosphorylation reactions. Therefore, insulin does not elicit an intracellular response.
The presence of more insulin will have no effect.

G proteins are molecular switches with two interconvertible forms, an active GTP-bound
form and an inactive GDP-bound form (Figure 9.42). In normal G proteins, GTPase activity
converts the active G protein to the inactive form. Because the ras protein lacks GTPase activity,

Chapter 10 SOLUTIONS

721

it cannot be inactivated. The result is continuous activation of adenylyl cyclase and prolonged
responses to certain extracellular signals.

17. The surface of a sphere is 47ir 2 . The surface area of the oocyte is 47r(50) 2 ^m, or 3.9 X 10 5 ^im 2 .
The surface area of a lipid molecule is 10 _14 cm 2 = lCT^m 2 . Since only 75% of the membrane
is lipid, the total number of lipid molecules is

3 9 X 1 0 5

— — X 0.75 = 2.9 X 10 11 molecules

10“ 6

18. Assuming that the lipid molecules made by your grandmother are equally divided between
daughter cells at each cell division, then after 30 cell divisions the oocyte (egg cell) produced
by your mother will have 1 / 2 30 of the original lipid molecules. Since the number of lipid mol-
ecules she inherited from her mother (your grandmother) was 2.9 X 10 11 (see previous
question), then the number remaining in each oocyte was

1/2 30 X 2.9 X 1 0 1 1 = 270
You inherited 270 lipid molecules from your grandmother.

Chapter 10 Introduction to Metabolism

1. (a)

(b) Inhibition of the first step in the common pathway by either G or J prevents the needless
accumulation of intermediates in the pathway. When there is ample G or J, fewer mole-
cules of A enter the pathway. By regulating an enzyme after the branch point, G or J
inhibits its own production without inhibiting production of the other.

2. Compartmentalizing metabolic processes allows optimal concentrations of substrates and
products for each pathway to exist independently in each compartment. In addition, separa-
tion of pathway enzymes also permits independent regulation of each pathway without inter-
ference by regulators from the other pathway.

3. Bacteria are much smaller than most eukaryotic cells so having separate compartments may
not be as much of an advantage. It’s also possible that localizing the citric acid cycle in mito-
chondria may be an historical accident rather than a selective advantage in eukaryotes.

4. In a multistep enzymatic pathway, the product from one enzyme will be the substrate for the
next enzyme in the pathway. For independent soluble enzymes, the product of each enzyme
must find the next enzyme by random diffusion in solution. By having sequential enzymes
located in close proximity to each other, either in a multienzyme complex or on a membrane,
the product of each enzyme can be passed directly on to the next enzyme without losing the
substrate by diffusion into solution.

5. (a) AG°' = RT In /C eq
AG°'

ln * eq = =
K eq = 38

-9000 I mol

i-i

(8.315 J K _l mol 1 )(298 K)

(b) AG°' = -RT In K eq

[Glucose][Pj]

K (0.1 M)(0.1 M)

eq [Glucose 6-P][H 2 0] (3.5 X 1(T 5 M)(1)

AG°' = -(8.315 JK” 1 mol“ 1 )(298 K) In 286
AG°' = -14 000) moP 1 = -14 kj mol 1
[Arginine][Pj]

= 3.63

= 286

6. (a) AG = AG°' + RT In- u . . in , ^

[Phosphoarginme][H 2 0]

AG = -32 000 J mol -1 + (8.315 J K _1 mol“ 1 )(298 K)ln
AG = -48 kj mol -1

(2.6 X 1 0~ 3 )(5 X 1 O' 3 )
(6.8 X 10“ 3 )(1)

722 SOLUTIONS Chapter 10

(b) A G°' is defined under standard conditions of 1 M concentrations of reactants and prod-
ucts. (The concentration of water is assigned a value of 1.) AG depends on the actual
concentrations of the reactants and products.

(c) Molecules with high free energies of hydrolysis, such as phosphoarginine and acetyl CoA,
are thermodynamically unstable but may be kinetically stable. These molecules are hy-
drolyzed very slowly in the absence of an appropriate catalyst.

AG°'(kJ mol -1 )

7. Glucose 1 -phosphate + UTP » UDP-glucose + PPj 0

PPi + H 2 0 > 2 Pi -29

AG°'-29

8. (a) Although ATP is rapidly utilized for energy purposes such as muscle contraction and

membrane transport, it is also rapidly resynthesized from ADP and Pj through interme-
diary metabolic routes. Energy for this process is supplied from the degradation of carbo-
hydrates, fats, and amino acids or from energy storage molecules such as muscle creatine
phosphate (CP + ADP — » ATP + C). With this rapid recycling, 50 grams total of ATP
and ADP is sufficient for the chemical energy needs of the body.

(b) The role of ATP is that of a free energy transmitter rather than an energy storage mole-
cule. As indicated in part (a), ATP is not stored, but is rapidly utilized in energy- requiring
reactions.

9. AG°' for the reaction of ATP and creatine is calculated as

AG°'(k] mol -1 )
+43

-32

+ 11

Creatine + Pj <—
ATP + H 2 Q < —
Creatine + ATP

-> Phosphocreatine + H 2 0

ADP + Pj

— > Phosphocreatine + ADP

The ratio of ATP to ADP needed to maintain a 20:1 ratio of phosphocreatine to creatine is
calculated from Equation 10.13. At equilibrium, AG = 0, so

[Phosphocreatine][ADP]

AG°' = —RT In

(20)[ADP]
n (1 )[ATP]
(20) [ADP]

[Creatine][ATP]

AG°' (11 000 J mol -1 )

RT (8.315 J K -1 mor 1 )(298 K)

= -4.44

(1 )[ATP]
[ATP]

= 1.2

i~2

[ADP]

= 1667:1

10 .

— C —
©NHo

O (Acyl adenylate)

tRNA
+AMP

H II

R— C— C — O — tRNA

®NH

11. AG°' = -RT\r\ K eq

[fructose-6-phosphate] 2
eq [glucose-6-phosphate] 1
AG°' = -(8.315 J K -1 mol _1 )(298 K) In 2
AG°' = -1.7 k] moP 1

Chapter 10 SOLUTIONS 723

12. (a) In K eq = —

A G°'
RT

(25 000 J mol" 1 )

(8.315 JK” 1 mol - )(298 K)

= - 10.1

Keq = 4.1 X 10“ 5

(b) A G°' for the coupled reaction is calculated as

A » B

ATP + H 2 Q < — > ADP + Pi

A + ATP + H 2 0
AC°'

B + ADP + P:

AC°'(kJ moP 1 )
+25
-32
-7

= 2.8

In K eq = -
K eq = 17

K eq for the coupled reaction is about 180,000 times larger than K eq in part (a).

[B][ADP][Pj] [B][ADP] [S]( 1)

e q

[fl

[A]

[A][ATP][H 2 0] [A] [ATP] [A] (400)

= 6800:1

Coupling the reaction to ATP hydrolysis increases the ratio of [B] to [A] by a factor of
about 166 million (6800 + (4.1 X 10 -5 ) = 1.6 X 10 8 ).

13. Electrons flow from the molecule with a more negative standard reduction potential to the
molecule with a more positive standard reduction potential.

(b) Succinate + Q — » Fumarate + QH 2

14. The standard reduction potentials in Table 10.4 refer to half- reactions that are written as
S ox + n e® — > S re d. Two half- reactions can be added to obtain the coupled oxidation-reduction
reaction by reversing the direction of the half-reaction involving the reduced species and revers-

ing the sign of its reduction potential.

QH 2 > Q + 2 H® + 2e®

AC°' = -nFAE°’ = -(2)(96.48 kj V -1 mol^XO.^V)
AC°' = -37 kj mol -1

+0.82

-0.03

£°'(V)

+0.23

-0.04

A E ot = 0.19 V

(b) y 2 0 2 + 2 H® + 2 e® > H 2 0

Succinate » Fumarate 2 H® + 2 e®

% 0 2 + Succinate > H 2 0 + Fumarate ^ ol = yg

A C°' = -(2)(96.48 kjV -1 mol“ 1 )(0.79V)

AC°' = -150 kj mol®

15. The expected results are as shown in the bottom graph. As NADH is formed in the reaction
mixture, the absorbance at 340 nm will increase (see Box 10.1).

+0 04

FADH 2 > FAD + 2 H® + 2 e®

Q + FADH 2

QH 2 + FAD

+0.22

A E°' = 0.26 V

Ar Arn , RT t [QH 2 ][FAD]

AT = AT n

nF [Q][FADH 2 ]

0.026 V (5

A E = 0.26 V In —

2 (1

1 0' 5 )(2 x 1 0 4 )
1 0 _4 )(5 x 10“ 3 )

A E = 0.26 V - 0.01 3(-3.9) = 0.31 V

AC = -nFAE = -(2)(96.48 kj V” 1 mor 1 )(0.31 V)

AC = -60 kj mol -1

724 SOLUTIONS Chapter 11

Theoretically, the oxidation of FADH 2 by ubiquinone liberates more than enough free
energy to drive ATP synthesis from ADP and Pj.

Chapter 1 1 Glycolysis

1. (a) 2 (see Figure 11.2 and Reaction 11.12)

(b) 2 (1 ATP is consumed by the fructokinase reaction, 1 ATP is consumed by the triose ki-
nase reaction, and 4 ATP are generated by the triose stage of glycolysis)

(d) 5 (2 ATP are obtained from fructose, as in part (b), and 3 ATP — rather than 2 — are ob-
tained from the glucose moiety since glucose 1 -phosphate, not glucose, is formed when
sucrose is cleaved)

H o

— n

’1

H — C — OH

2 |

3 coo©

H — C— OH

2 |

HO — C — H

3 1

Glycolysis +

H — C— OH

4 COO©

H — C— OH

5 |

HO — c — H

b l

6 ch 2 oh

6 ch 3

Glucose 2 Lactate

(b) Glucose labeled at either C-3 or C-4 yields 14 CQ 2 from the decarboxylation of pyruvate.

H ,0

’i

H — C — OH

HO — C — H

3 |

H — C— OH

-OH

6 CH 2 OH

Glucose

H o

(3,4) |

S — CoA
1

B „c=o

„=,c=o

(,„ CH 3

„6,CH 3

(2) Glyceraldehyde

(2) Pyruvate

(2) Acetyl CoA

3-phosphate

3. Inorganic phosphate ( 32 Pi) will be incorporated into 1,3-frisphosphoglycerate (1,3 BPG) at the
C-l carbon in the glyceraldehyde 3-phosphate dehydrogenase (GADPH) reaction — glycer-
aldehyde 3 -phosphate + NAD® + Pj — > 1,3 BPG — and then transferred to the y-position
of ATP in the next step: 1,3 BPG + ADP — » ATP + 3-phosphoglycerate.

4. Since the brain relies almost solely on glucose for energy, it is dependent on glycolysis as the
major pathway for glucose catabolism. Since the Huntington protein binds tightly to
GAPDH, this suggests that it might inhibit this crucial glycolytic enzyme and thereby impair
the production of ATP. Decreased ATP levels would be detrimental to neuronal cells in the
brain.

ch 2 oh

ATP

ADP

CH 2 OH

NAD©

ch 2 oh

NADH, H® U

— C — H

s ,

HO — C — H

. r=n

2 i

ch 2 oh

3 Z

Glycerol

2| ©

ch 2 opo 3 (l)

L-Glycerol

3-phosphate

2 i ©

CHjOPOj^

Dihydroxyacetone

phosphate

(b) C-2 and C-3 of glycerol 3-phosphate must be labeled. Once dihydroxyacetone phosphate
is converted to glyceraldehyde 3-phosphate, C-l is oxidized to an aldehyde and subse-
quently lost as C0 2 (Problem 2).

6. Cells that metabolize glucose to lactate by anaerobic glycolysis produce far less ATP per glu-
cose than do cells that metabolize glucose aerobically to C0 2 via glycolysis and the citric acid
cycle (Figure 11.1). More glucose must be utilized via anaerobic glycolysis to produce a suffi-
cient amount of ATP for cellular needs, and the rate of conversion of glucose to lactate is

Chapter 12 SOLUTIONS

725

much higher than under aerobic conditions. Cancer cells in an anaerobic environment take
up far more glucose and may overproduce some glycolytic enzymes to compensate for the in-
crease in the activity of this pathway of carbohydrate metabolism.

7. No. The conversion of pyruvate to lactate, catalyzed by lactate dehydrogenase, oxidizes
NADH to NAD©, which is required for the glyceraldehyde 3 -phosphate dehydrogenase reac-
tion of glycolysis.

8. In the reactions catalyzed by these enzymes, the bond between the y-phosphorus atom and
the oxygen of the /3-phosphoryl group is cleaved when the y-phosphoryl group of ATP is
transferred (Figure 11.3). The analog cannot be cleaved in this way and therefore inhibits the
enzymes by competing with ATP for the active site.

9 . The free energy change for the aldolase reaction under standard conditions (A G°') is
+22.8 kj mol -1 . The concentrations of fructose 1,6-Hsphosphate, dihydroxyacetone phos-
phate, and glyceraldehyde 3-phosphate in heart muscle, however, are much different than the
1 M concentrations assumed under standard conditions. The actual free energy change under
cellular concentrations (A G°' = — 5.9 kj mol -1 ) is much different than A G°', and the al-
dolase reaction readily proceeds in the direction necessary for glycolysis: Fructose
1, 6-frisphosphate —> glyceraldehyde 3-phosphate + dihydroxyacetone phosphate.

10. The standard Gibbs free energy change is + 28 kj mol -1 . The equilibrium constant is

11 .

(a)

(b)

28 = RT In K f

[DHAP][G3P] = [5^

[FBP]

250/ulM

25,000 ,uM = 25 mM

10“ 6 ][5 :
[FBP]

1 0 5 (Equation 1.12)

10 -6 ]

= 10" 5 FBP = 2.5

/xM

ATP is both a substrate and an allosteric inhibitor for PFK- 1 . Higher concentrations of
ATP result in a decrease in the activity of PFK- 1 due to an increase in the K m . AMP is an
allosteric activator that acts by relieving the inhibition caused by ATP, thus raising the
curve when AMP is present with ATP.

F2,6P is an allosteric activator of PFK-1. In the presence of F2,6P the activity of PFK-1 is
increased due to a decrease in the apparent K m for fructose 6-phosphate.

12 . Increased [cAMP] activates protein kinase A, which catalyzes the phosphorylation and inac-
tivation of pyruvate kinase.

cAMP

Pyruvate kinase
(more active)

Protein kinase A

7 \ *

13 . (a) A decrease in glycolysis in the liver makes more glucose available for export to other

tissues.

(b) Decreased activity of the glucagon transducer system decreases the amount of cAMP
formed. As existing cAMP is hydrolyzed by the activity of a phosphodiesterase, cAMP-
dependent protein kinase A becomes less active. Under these conditions, PFK-2 activity
increases and fructose 2,6-frisphosphatase activity decreases (Figure 11.18). The resulting
increase in fructose 2,6-frisphosphate activates PFK-1, increasing the overall rate of gly-
colysis. A decrease in cAMP also leads to the activation of pyruvate kinase (Problem 12).

14 . Chemoautotrophs use glycolysis to generate energy from stored glucose residues in glycogen
as described in Chapter 12.

Chapter 12 Gluconeogenesis, The Pentose Phosphate Pathway,
and Glycogen Metabolism

1. 2 pyruvate + 2NADH + 4 ATP + 2 GTP + 6 H 2 0 + 2 H© ^
glucose + 2 NAD© + 4 ADP + 2 GDP + 6 Pj
2 NADH = 5 ATP equivalents

4 ATP =4 ATP
2 ATP

2GTP

11 ATP

726

SOLUTIONS Chapter 12

The energy required to synthesize one molecule of glucose 6-phosphate from C0 2 can be cal-
culated from Reaction 12.7.

12 NADPH = 30 ATP

The conversion of G6P to glucose does not require or produce ATP equivalents. The synthe-
sis of glucose from pyruvate via the gluconeogenesis pathway is only about one third (11/30)
as expensive as the synthesis of glucose from C0 2 .

2. Reducing power in the form of NADH (2), and ATP (4) and GTP (2) are required for the syn-
thesis of glucose from pyruvate (Equation 12.1). The NADH and GTP are direct products of
the citric acid cycle, and ATP can be generated from NADH and QH 2 (FADH 2 ) during the ox-
idative phosphorylation process.

3. Epinephrine interacts with the liver /3 -adrenergic receptors and activates the adenylyl cyclase
signaling pathway, leading to cAMP production and activation of protein kinase A (Figure
12.15). Protein kinase A activates phosphorylase kinase, which in turn activates glycogen
phosphorylase (GP), leading to glycogen degradation (Figure 12.16). Glucose can then be
transported out of the liver and into the bloodstream, where it is taken up by muscles for
needed energy production.

(Gp\

Liver[Glycogen — — GIP » G6P » Glucose] » Bloodstream » Muscles

4. (a) Protein phosphatase- 1 activated by insulin catalyzes the hydrolysis of the phosphate ester

bonds on glycogen synthase (activating it) and on glycogen phosphorylase and phospho-
rylase kinase (inactivating them), as shown in Figure 12.17. Therefore, insulin stimulates
glycogen synthesis and inhibits glycogen degradation in muscle cells.

(b) Only liver cells are rich in glucagon receptors, so glucagon selectively exerts its effects on
liver enzymes.

(c) The binding of glucose to the glycogen phosphorylase-protein phosphatase- 1 complex
in liver cells relieves the inhibition of protein phosphatase- 1 and makes glycogen phos-
phorylase more susceptible to dephosphorylation (inactivation) by protein phosphatase- 1
(Figure 12.18). Protein phosphatase- 1 also catalyzes the dephosphorylation of glycogen
synthase, making it more active. Therefore, glucose stimulates glycogen synthesis and in-
hibits glycogen degradation in the liver.

5. Decreased concentrations of fructose 2,6-frzsphosphate (F2,6BP) lead to a decreased rate of
glycolysis and an increased rate of gluconeogenesis. F2,6BP is an activator of the glycolytic
enzyme phosphofructokinase- 1 (PFK-1), and lower F2,6BP levels will result in decreased
rates of glycolysis. In addition, F2,6BP is an inhibitor of the gluconeogenic enzyme fructose
1,6-Znsphosphatase, and therefore decreased levels of F2,6BP will decrease the inhibition and
increase the rate of gluconeogenesis (Figure 12.4).

6. When glucagon binds to its receptor, it activates adenylyl cyclase. Adenylyl cyclase catalyzes
the synthesis of cAMP from ATP. The cAMP activates protein kinase A. Protein kinase A cat-
alyzes the phosphorylation of PFK-2, which inactivates the kinase activity and activates the
phosphatase activity. Fructose 2,6-Znsphosphatase catalyzes the hydrolytic dephosphorylation
of fructose 2,6-Msphosphate to form fructose 6-phosphate. The resulting decrease in the con-
centration of fructose 2,6-/7zsphosphate relieves the inhibition of fructose 1,6-frisphosphatase,
thereby activating gluconeogenesis. Thus, the kinase activity of PFK-2 is decreased.

7. (a) Yes. The synthesis of glycogen from glucose 6-phosphate requires the energy of one phos-

phoanhydride bond (in the hydrolysis of PPj; Figure 12.10). However, when glycogen is
degraded to glucose 6-phosphate, inorganic phosphate (Pj) is used in the phosphorolysis
reaction. No “high energy” phosphate bond is used.

(b) One fewer ATP molecule is available for use in the muscle when liver glycogen is the source
of the glucose utilized. Fiver glycogen is degraded to glucose phosphates and then to glu-
cose without consuming ATP. After transport to muscle cells, the glucose is converted to
glucose 6-phosphate by the action of hexokinase in a reaction that consumes one molecule
of ATP. Muscle glycogen, however, is converted directly to glucose 1 -phosphate by the
action of glycogen phosphorylase, which does not consume ATP. Glucose 1 -phosphate is
isomerized to glucose 6 -phosphate by the action of phosphoglucomutase.

8. A deficiency of glycogen phosphorylase in the muscle prevents the mobilization of glycogen
to glucose. Insufficient glucose prevents the production of ATP by glycolysis. Existing ATP
used for muscle contraction is not replenished, thus increasing the levels of ADP and Pj. Since
no glucose is available from glycogen in the muscle, no lactate is produced.

9. Converting glucose 1 -phosphate to two molecules of lactate yields 3 ATP equivalents (1 ATP
expended in the phosphofructokinase- 1 reaction, 2 ATP produced in the phosphoglycerate

Chapter 13 SOLUTIONS

727

kinase reaction, and 2 ATP produced in the pyruvate kinase reaction). Converting two mole-
cules of lactate to one molecule of glucose 1 -phosphate requires 6 ATP equivalents (2 ATP in
the pyruvate carboxylase reaction, 2 GTP in the PEP carboxykinase reaction, and 2 ATP in the
phosphoglycerate kinase reaction).

10. (a) Muscle pyruvate from glycolysis or amino acid catabolism is converted to alanine by

transamination. Alanine travels to the liver, where it is reconverted to pyruvate by
transamination with a-ketoglutarate. Gluconeogenesis converts pyruvate to glucose,
which can be returned to muscles.

(b) NADH is required to reduce pyruvate to lactate in the Cori cycle, but it is not required to
convert pyruvate to alanine in the glucose- alanine cycle. Thus, the glucose-alanine cycle
makes more NADH available in muscles for the production of ATP by oxidative phos-
phorylation.

11. (a) Inadequate glucose 6-phosphatase activity (G6P —> glucose + Pj) leads to accumulation

of intracellular G6P, which inhibits glycogen phosphorylase and activates glycogen syn-
thase. This prevents liver glycogen from being mobilized. This results in increased glyco-
gen storage (and enlargement of the liver) and low blood glucose levels (hypoglycemia).

(b) Yes. A defective branching enzyme leads to accumulation of glycogen molecules with de-
fective, short outer branches. These molecules cannot be degraded, so there will be much
less efficient glycogen degradation for glucose formation. Low blood glucose levels result
due to the impaired glycogen degradation.

(c) Inadequate liver phosphorylase activity leads to an accumulation of liver glycogen since
the enzyme cleaves a glucose molecule from the nonreducing end of a glycogen chain.

Low blood glucose levels result, due to the impaired degradation of glycogen.

12. Glucose 6-phosphate, glyceraldehyde 3-phosphate, and fructose 6-phosphate.

13. The repair of tissue injury requires cell proliferation and synthesis of scar tissue. NADPH is
needed for the synthesis of cholesterol and fatty acids (components of cellular membranes),
and ribose 5-phosphate is needed for the synthesis of DNA and RNA. Since the pentose phos-
phate pathway is the primary source of NADPH and ribose 5-phosphate, injured tissue re-
sponds to the increased demands for these products by increasing the level of synthesis of the
enzymes in the pentose phosphate pathway.

14. (a) CH 2 OH

ch 2 oh

C =0
1

HO — C — H

H — C — OH

L = U

HO — C — H

H — C — OH

| +

Transketolase

H — C — OH

CH 2 0P0 3 ©

ch 2 opo|^

ch 2 opo 3 ©

Xylulose 5-phosphate Erythrose 4-phosphate Glyceraldehyde 3-phosphate Fructose 6-phosphate

(b) C-2 of glucose 6-phosphate becomes C-l of xylulose 5-phosphate. After C-l and C-2 of
xylulose 5-phosphate are transferred to erythrose 4-phosphate, the label appears at C-l
of fructose 6-phosphate, as shown in part (a).

Chapter 13 The Citric Acid Cycle

1. (a) No net synthesis is possible since two carbons from acetyl Co A enter the cycle in the cit-

rate synthase reaction and two carbons leave as C0 2 in the isocitrate dehydrogenase and
a-ketoglutarate dehydrogenase reactions.

(b) Oxaloacetate can be replenished by the pyruvate carboxylase reaction, which carries out a
net synthesis of OAA,

Pyruvate + C0 2 + ATP + H 2 0 > Oxaloacetate + ADP + Pj

This is the major anaplerotic reaction in some mammalian tissues. Many plants and
some bacteria supply oxaloacetate via the phosphoenolpyruvate carboxykinase reaction,

Phosphoenolpyruvate + HCO^ » Oxaloacetate + Pj

In most species, acetyl CoA can be converted to malate and oxaloacetate via the glyoxylate
pathway.

2. Aconitase would be inhibited by fluorocitrate formed from fluoroacetate, leading to increased
levels of citric acid and decreased levels of all subsequent citric acid cycle intermediates from

728

SOLUTIONS Chapter 13

isocitrate to oxaloacetate. Since fluorocitrate is a competitive inhibitor, very high levels of cit-
rate would at least partially overcome the inhibition of aconitase by fluorocitrate and permit
the cycle to continue at some level.

3. (a) 12.5; 10.0 from the cycle and 2.5 from the pyruvate dehydrogenase reaction.

(b) 10.0; 7.5 from oxidation of 3 NADH, 1.5 from oxidation of 1 QH 2 , and 1.0 from the
substrate-level phosphorylation catalyzed by CoA synthetase.

4. 87.5% (28 of 32) of the ATP is produced by oxidative phosphorylation, and 12.5% (4 of 32) is
produced by substrate-level phosphorylation.

5. Thiamine is the precursor of the coenzyme thiamine pyrophosphate (TPP) , which is found in
two enzyme complexes associated with the citric acid cycle: the pyruvate dehydrogenase com-
plex and the a-ketoglutarate dehydrogenase complex. A deficiency of TPP decreases the ac-
tivities of these enzyme complexes. Decreasing the conversion of pyruvate to acetyl CoA and
of a-ketoglutarate to succinyl CoA causes accumulation of pyruvate and a-ketoglutarate.

6. Since C- 1 of pyruvate is converted to C0 2 in the reaction catalyzed by the pyruvate dehydro-
genase complex, 1-[ 14 C] -pyruvate is the first to yield 14 C0 2 . Neither of the two acetyl carbon
atoms of acetyl CoA is converted to C0 2 during the first turn of the citric acid cycle (Figure
13.5). However, the carboxylate carbon atoms of oxaloacetate, which arise from C-2 of pyru-
vate, become the two carboxylates of citrate that are removed as C0 2 during a second turn of
the cycle. Therefore, 2- [ 14 C] -pyruvate is the second labeled molecule to yield
14 C0 2 . 3- [ 14 C] -Pyruvate is the last to yield 14 C0 2 , in the third turn of the cycle.

First turn

coo°

,C0 2

S-CoA

,c=o

7 7

h°

ch 3

3 ch 3

Pyruvate Acetyl CoA

2 COO°

3 ch 2

C0 2 C0 2

2 COO°

— c— coo®
1

7 7 ,

3 ch 2

cn.

3 ch 2

coo u

2 COO^

Citrate

Succinate

Second turn

2 COO°

coo°

3 C00°

ChH,

2 C0 2

(~ r\ :

v MA r

7 ,

3 ch 2

3 I

3 ch 2

1 ©

3 COO u

1 0

2 COO u

1 0

2 COO^

Oxaloacetate

Citrate

Succinate

Half of the 14 C is eliminated by the third turn of the cycle. An additional one-fourth is elimi-
nated in the fourth turn, then one-eighth in the fifth turn, etc. It will take a very long time to
eliminate all of the 14 C from the citric acid cycle intermediates.

7. (a) The NADH produced by the oxidative reactions of the citric acid cycle must be recycled

back to NAD®, which is required for the pyruvate dehydrogenase reaction. When 0 2 levels
are low, fewer NADH molecules are reoxidized by 0 2 (via the process of oxidative phos-
phorylation), so the activity of the pyruvate dehydrogenase complex decreases.

(b) Pyruvate dehydrogenase kinase catalyzes phosphorylation of the pyruvate dehydrogenase
complex, thereby inactivating it (Figure 13.12). Inhibiting the kinase shifts the pyruvate
dehydrogenase complex to its more active form.

8. A deficiency in the citric acid cycle enzyme fumarase would result in abnormally high concen-
trations of fumarate and prior cycle intermediates including succinate and a-ketoglutarate,
which could lead to excretion of these molecules.

9. The different actions of acetyl CoA on two components of the pyruvate dehydrogenase
(PDH) complex both lead to an inhibition of the pyruvate to acetyl CoA reaction. Acetyl CoA
inhibits the E 2 component of the PDH complex directly (Figure 13. 1 1). Acetyl CoA causes in-
hibition of the Ei component indirectly by activating the pyruvate kinase (PK) component of

Chapter 13 SOLUTIONS

729

the PDH complex, and PK phosphorylates the E : component of the PDH complex, thus inac-
tivating it (Figure 13.12).

10. The pyruvate dehydrogenase complex catalyzes the oxidation of pyruvate to form acetyl CoA
and C0 2 . If there is reduced activity of this complex, then the pyruvate concentration will in-
crease. Pyruvate will be converted to lactate through the action of lactate dehydrogenase. Lac-
tate builds up since glycolytic metabolism is increased to synthesize ATP since oxidation of
pyruvate to acetyl CoA is impaired. In addition, pyruvate is converted to alanine, as shown in
Reaction 12.6.

11. Calcium activates both isocitrate dehydrogenase and a-ketoglutarate dehydrogenase in the
citric acid cycle, thereby increasing this catabolic process and producing more ATP. In addi-
tion, Ca© activates the pyruvate dehydrogenase phosphatase enzyme of the PDH complex,
which activates the Ei component (Figure 13.12). Activation of the PDH complex converts
more pyruvate into acetyl CoA for entry into the citric acid cycle, resulting in an increased
production of ATP.

12. (a) Alanine degradation replenishes citric acid cycle intermediates, since pyruvate can be
converted to oxaloacetate via the pyruvate carboxylase reaction, the major anaplerotic re-
action in mammals (Reaction 13.19). Leucine degradation cannot replenish intermedi-
ates of the citric acid cycle, since for every molecule of acetyl CoA that enters the cycle,
two molecules of C0 2 are lost.

(b) By activating pyruvate carboxylase, acetyl CoA increases the amount of oxaloacetate pro-
duced directly from pyruvate. The oxaloacetate can react with the acetyl CoA produced
by the degradation of fatty acids. As a result, flux through the citric acid cycle increases to
recover the energy stored in the fatty acids.

13. (a)

COO©

l ” 2

»ch 2

c=o

coo©

-Ketoglutarate

(b)

Ala

C H 3

o=c — coo©

(Pyruvate)

^CCb

( c ) 14 coo e
I

ch 2

0 = C — COO e
Oxaloacetate

C h 3

0= 4 C — SCoA

(Acetyl SCoA)

ch 2 coo©

HO — C — COO 0 Citrate
CH 22 — COO 0

14. (a) Two molecules of acetyl CoA yield 20 ATP molecules via the citric acid cycle (Figure

13.10) or 6.5 ATP molecules via the glyoxylate cycle (from the oxidation of two molecules
of NADH and one molecule of QH 2 ; Reaction 13.22).

(b) The primary function of the citric acid cycle is to oxidize acetyl CoA to provide the re-
duced coenzymes necessary for the generation of energy- rich molecules such as ATP. The
primary function of the glyoxylate cycle is not to produce ATP, but to convert acetyl
groups to four-carbon molecules that can be used to produce glucose.

15. The protein that controls the activity of isocitrate dehydrogenase in E. coli is a bifunctional
enzyme with kinase and phosphatase activities in the same protein molecule. The kinase ac-
tivity phosphorylates isocitrate dehydrogenase to inhibit the activity of isocitrate dehydro-
genase, and the phosphatase activity dephosphorylates isocitrate dehydrogenase to activate
isocitrate dehydrogenase. When concentrations of glycolytic and citric acid cycle intermedi-
ates are high, isocitrate dehydrogenase is not phosphorylated and is active. When phospho-
rylation decreases the activity of isocitrate dehydrogenase, isocitrate is diverted to the
glyoxylate cycle.

730

SOLUTIONS Chapter 14

Chapter 14 Electron Transport and Oxidative Phosphorylation

1. The formula for calculating protonmotive force is

AC = F Aip - 2.303 RT A pH
If C = -21 ,000 kj and At// = -0.1 5 V, then at 25°C

-21,200 = (96485 X -0.15) - 2.303(8.315 X 298) ApH
5707 A pH = 6727
A pH = 1.2

Since the outside pH is 6.35 and the inside is negative (higher pH), then the cytoplasmic pH
is 6.35 + 1.2 = 7.55.

2. The reduction potential of an iron atom in a heme group depends on the surrounding pro-
tein environment, which differs for each cytochrome. The differences in reduction potentials
allow electrons to pass through a series of cytochromes.

3. Refer to Figure 14.6.

(a) Complex III. The absence of cytochrome c prevents further electron flow.

(b) No reaction occurs since Complex I, which accepts electrons from NADH, is missing.

(d) Cytochrome c. The absence of Complex IV prevents further electron flow.

4. UCP-2 leaks protons back into the mitochondria, thereby decreasing the protonmotive force.
The metabolism of foodstuffs provides the energy for electron transport, which in turn cre-
ates the protonmotive gradient used to produce ATR An increase in UCP-2 levels would make
the tissue less metabolically efficient (i.e., less ATP would be produced per gram of foodstuff
metabolized). As a result, more carbohydrates, fats, and proteins would have to be metabo-
lized in order to satisfy the basic metabolic needs, and this could “burn off” more calories and
potentially cause weight loss.

5. (a) Demerol interacts with Complex I and prevents electron transfer from NADH to Q. The

concentration of NADH increases since it cannot be reoxidized to NAD®. The concen-
tration of Q increases since electrons from QH 2 are transferred to 0 2 but Q is not re-
duced back to QH 2 .

(b) Myxothiazole inhibits electron transfer from QH 2 to cytochrome q and from QH 2 (via • Q - )
to cytochrome b 566 in Complex III (Figure 14.14). The oxidized forms of both cytochromes
predominate since Fe© cannot be reduced by electrons from QH 2 .

14.19), and it is prevented from doing so by the binding of CN® to the iron atom.

(b) The methemoglobin (Fe©) generated from nitrite treatment competes with cytochrome
a 3 for the CN® ions. This competition effectively lowers the concentration of cyanide
available to inhibit cytochrome a 3 in Complex IV, and decreases the inhibition of the
electron transport chains in the presence of CN®.

7. A substrate is usually oxidized by a compound with a more positive reduction potential. Since
E°' for the fatty acid is close to E° f for FAD in Complex II (0.0 V, as shown in Table 14. 1), elec-
tron transfer from the fatty acid to FAD is energetically favorable.

A E°' = 0.0 V - (-0.05 V) = +0.05 V

AC°' = -nFAE 0 '

AC°' = -(2)(96.48 kj V _1 )(0.05 V) = -9.6 kj mol -1

Since E°' for NADH in Complex I is —0.32 V, the transfer of electrons from the fatty acid to
NADH is unfavorable.

A P' = -0.32 V - (-0.05 V) = -0.27 V

AC°' = -(2)(96.48 kj NT 1 mor’X-O^? V) = 52 kj mcT 1

8. (a) 10 protons; 2.5 ATP; P : O = 2.5.

(b) 6 protons; 1.5ATP;P:0 = 1.5.

9. (a) The inner mitochondrial membrane has a net positive charge on the cytosolic side (out-

Chapter 15 SOLUTIONS

731

net movement of one negative charge from the inner matrix side to the positive cytosolic
side. The membrane potential thereby assures that outward transport of a negatively
charged ATP is favored by the outside positive charge.

(b) Yes. The electrochemical potential with a net positive charge outside the membrane is a
result of proton pumping, which is driven by the electron transport chain. This in turn
requires oxidation of metabolites to generate NADH and QH 2 as electron donors.

10. ATP synthesis is normally associated with electron transport. Unless ADP can continue to be
translocated into the mitochondrial matrix for the ATP synthesis reaction (ADP + Pj —> ATP),
ATP synthesis will not occur and the proton gradient will not be dissipated. Electron transport
will be inhibited as the proton concentration increases in the intermembrane space.

11. (a) AC = PAT' - 2.303 RT A pH (Equation 14.6)

AC = ((96485)(-0.18)) - ((2.303)(8.31 5)(0.7))

AC = -17367 - 3995
AC = -2136 = 21 kj mol -1

(b) AG tota i = 21.36 kj mol -1

Charge gradient contribution is 17.367 kj mol -1 , or 17.367 ^ 21.36 X 100 = 81.3%
pH gradient contribution is 3.995 kj mol -1 , or 3.995 -r- 21.36 X 10 = 18.7%

12. (a) In the malate-aspartate shuttle, the reduction of oxaloacetate in the cytosol consumes a

proton that is released in the matrix by the oxidation of malate (Figure 14.27). Therefore,
one fewer proton is contributed to the proton concentration gradient for every cytosolic
NADH oxidized (9 versus 10 for mitochondrial NADH). The ATP yield from two mole-
cules of cytoplasmic NADH is about 4.5 rather than 5.0.

(b) Cytoplasmic reactions

Glucose > 2 Pyruvate 2.0 ATP

2 NADH » 4.5 ATP

Mitochondrial reactions

2 Pyruvate » 2 Acetyl CoA + 2 C0 2 2 NADH » 5.0 ATP

2 Acetyl CoA > 4 C0 2 2.0 GTP

6 NADH > 15.0 ATP

2 QH 2 > 3.0 ATP

Total 31.5 ATP

Chapter 15 Photosynthesis

1. Because in photosynthesis there are two steps where light energy is absorbed to produce “high
energy” electrons, thus PS II transfers 6H® insead of 10 H® in respiration but PSI produces
2.5 ATP equivalents — the same as respiration.

2. Plant chlorophylls absorb energy in the red region of the spectrum (Figure 15.2). The
dragonfish chlorophyll derivatives absorb the red light energy (667 nm), and pass the sig-
nals on to the visual pigments in much the same manner that plant antenna chlorophylls
and related molecules capture light energy and transfer it to a reaction center where elec-
trons are promoted into excited states for transfer to acceptors of the electron transport
chain.

3. (a) Rubisco is the world s most abundant protein and the principal catalyst for photosynthesis,

the basic means by which living organisms acquire the carbon necessary for life. Its im-
portance in the process of providing food for all living things can be well justified.

(b) Photorespiration is a process that wastes ribulose 1,5-frisphosphate, consumes the
NADPH and ATP generated by the light reactions, and can greatly reduce crop yields. As
much as 20% to 30% of the carbon fixed in photosynthesis can be lost to photo respira-
tion. This process results from the lack of specificity of Rubisco, which can use 0 2 instead
of C0 2 (Figure 15.8) to produce phosphoglycolate and 3-phosphoglycerate (Figure
15.18) instead of two triose phosphate molecules. In addition, Rubisco has low catalytic
activity (K cat ~ 3 s _1 ). This lack of specificity and low activity earns Rubisco the title of a
relatively incompetent, inefficient enzyme.

4. 6C0 2 + 6H 2 S^C 6 H 12 0 6 + 3 0 2 + 6 S
6 C0 2 + 12 H® — » C 6 H 12 Og + 3 0 2

732

SOLUTIONS Chapter 15

5. (a) C0 2 + 2 H 2 S y > (CH 2 0) + H 2 0 + 2 S

C0 2 + 2 CH 3 CH 2 OH Ught > (CH 2 0) + H 2 0 + 2 CH 3 CHO

Ethanol Acetaldehyde

(b) When H 2 0 is the proton donor, 0 2 is the product, but when other proton donors such as
H 2 S and ethanol are used, oxygen cannot be produced. Most photosynthetic bacteria do
not produce 0 2 and are obligate anaerobes that are poisoned by 0 2 .

6. Rubisco is not active in the dark because it requires alkaline conditions. Those conditions
only occur when photosynthesis is active so there’s nothing (except light) that can be added to
the chloroplast suspension in the dark that will activate the calvin cycle.

7. (a) Two H 2 0 molecules provide the oxygens for one 0 2 during the photosynthetic process. A

total of four electrons must be removed from two H 2 0 and passed through an electron
transport system to two NADPH. One quantum of light is required to transfer one elec-
tron through PSI and one quantum for PSII. Therefore, a total of eight photons will be re-
quired to move four electrons through both reaction centers (four photons for PHI and
four photons for PHII).

(b) Six NADPH are required for the synthesis of one triose phosphate by the Calvin cycle
(Figure 15.21). Therefore, 12 electrons must be transferred through the two reaction cen-
ters of the electron transport system and this will require the absorption of 24 hv.

8. (a) Yes. (Refer to the Z-scheme, Figure 15.14). When DCMU blocks electron flow, PSII in the

P680* state will not be reoxidized to the P680© state, which is required as an acceptor of
electrons from H 2 0. If H 2 0 is not oxidized by P680®, then no 0 2 will be produced. In
the absence of electron flow through the cytochrome bf complex, no protons will be
translocated across the membrane. Without a pH gradient no photophosphorylation
(ATP synthesis) will be possible.

(b) External electron acceptors for PSII will permit P680 to be reoxidized to P680© and will
restore 0 2 evolution. No electrons will flow through the cytochrome bf complex, how-
ever, so no photophosphorylation will occur.

9. (a) When the external pH rises to 8.0, the stromal pH also rises quickly, but the luminal pH

remains low initially because the thylakoid membrane is relatively impermeable to pro-
tons. The pH gradient across the thylakoid membrane drives the production of ATP via
proton translocation through chloroplast ATP synthase (Figure 15.16).

(b) Protons are transferred from the lumen to the stroma by ATP synthase, driving ATP syn-
thesis. The pH gradient across the membrane decreases until it is insufficient to drive the
phosphorylation of ADP, and ATP synthesis stops.

10. During cyclic electron transport, reduced ferredoxin donates its electrons back to P700 via
the cytochrome bf complex (Figure 15.11). As these electrons cycle again through photosys-
tem I, the proton concentration gradient generated by the cytochrome bf complex drives ATP
synthesis. However, no NADPH is produced because there is no net flow of electrons from
H 2 0 to ferredoxin. No 0 2 is produced because photosystem II, the site of 0 2 production, is
not involved in cyclic electron transport.

11. The light absorbing complexes, electron transport chain, and chloroplast ATP synthase all reside
in the thylakoid membranes, and the structure and interactions of any of these photosynthetic
components could be affected by a change in the physical nature of the membrane lipids.

12. The compound is acting as an uncoupler. The electron transfer is occurring without the
synthesis of ATP. The compound destroys the proton gradient that is produced through
electron transfer.

13. (a) The synthesis of one triose phosphate from C0 2 requires 9 molecules of ATP and 6 mole-

cules of NADPH (Equation 15.5). Since two molecules of triose phosphate can be converted
to glucose, glucose synthesis requires 18 molecules of ATP and 12 molecules of NADPH.
(b) Incorporating glucose 1 -phosphate into starch requires one ATP equivalent during the
conversion of glucose 1 -phosphate to ADP-glucose (Figure 15.24), bringing the total re-
quirement to 19 molecules of ATP and 12 molecules of NADPH.

14. Refer to Figure 15.21. (a) C-l. (b) C-3 and C-4. (c) C-l and C-2. C-l and C-2 of fructose
6-phosphate are transferred to glyceraldehyde 3-phosphate to form xylulose 5-phosphate. C-3
and C-4 of fructose 6-phosphate become C-l and C-2 of erythrose 4-phosphate.

15. (a) In the C 4 pathway (Figure 15.29), the pyruvate-phosphate dikinase reaction consumes

two ATP equivalents for each C0 2 fixed (since PPi is hydrolyzed to 2 Pj). Therefore, C 4

Chapter 16 SOLUTIONS

733

plants require 12 more molecules of ATP per molecule of glucose synthesized than C 3
plants require.

(b) Because C 4 plants minimize photorespiration, they are more efficient than C 3 plants in
using light energy to fix C0 2 into carbohydrates, even though the chemical reactions for
fixing C0 2 in C 4 plants require more ATP.

16. (a) An increase in stromal pH increases the rate of the Calvin cycle in two ways.

(1) An increase in stromal pH increases the activity of ribulose l,5-/7zsphosphate carboxy-
lase-oxygenase (Rubisco), the central regulatory enzyme of the Calvin cycle, and the
activities of fructose 1,6-frzsphosphatase and sedoheptulose l,7-/7zsphosphatase. It also
increases the activity of phosphoribulokinase. Phosphoribulokinase is inhibited by 3-
phosphoglycerate (3PG) in the 3PG© ionization state but not in the 3PG© ioniza-
tion state, which predominates at higher pH.

(2) An increase in stromal pH also increases the proton gradient that drives the synthesis
of ATP in chloroplasts. Since the reactions of the Calvin cycle are driven by ATP, an
increase in ATP production increases the rate of the Calvin cycle.

(b) A decrease in the stromal concentration of Mg© decreases the rate of the Calvin cycle by
decreasing the activity of Rubisco, fructose 1,6-frisphosphatase, and sedoheptulose 1,7-
frisphosphatase.

Chapter 16 Lipid Metabolism

1. (a) LDLs are rich in cholesterol and cholesterol esters and transport these lipids to peripheral

tissues. Delivery of cholesterol to tissues is moderated by LDL receptors on the cell mem-
branes. When LDL receptors are defective, receptor- mediated uptake of cholesterol does
not occur (Section 16.1 OB). Because cholesterol is not cleared from the blood it accumu-
lates and contributes to the formation of atherosclerotic plaques.

(b) Increased cholesterol levels normally repress transcription of HMG-CoA reductase and
stimulate the proteolysis of this enzyme as well. With defective LDL, however, cholesterol
synthesis continues in spite of high blood cholesterol levels because the extracellular cho-
lesterol cannot enter the cells to regulate intracellular synthesis.

(c) HDLs remove cholesterol from plasma and cells of nonhepatic tissues and transport it to
the liver where it can be converted into bile salts for disposal. In Tangier patients, defec-
tive cholesterol-poor HDLs cannot absorb cholesterol, and the normal transport process
to the liver is disrupted.

2. (a) Carnitine is required to transport fatty acyl Co A into the mitochondrial matrix for

f3 -oxidation (Figure 16.24). The inhibition of fatty acid transport caused by a deficiency
in carnitine diminishes energy production from fats for muscular work. Excess fatty acyl
Co A can be converted to triacylglycerols in the muscle cells.

(b) Since carnitine is not required to transport pyruvate, a product of glycolysis, into mito-
chondria for oxidation, muscle glycogen metabolism is not affected in individuals with a
carnitine deficiency.

3. (a) Activation of the C 12 fatty acid to a fatty acyl Co A consumes 2 ATR Five rounds of

(3 -oxidation generate 6 acetyl CoA, 5 QH 2 (which yield 7.5 ATP via oxidative phosphory-
lation), and 5 NADH (which yield 12.5 ATP). Oxidation of the 6 acetyl CoA by the citric
acid cycle yields 60 ATP. Therefore, the net yield is 78 ATP equivalents.

(b) Activation of the C 16 monounsaturated fatty acid to a fatty acyl CoA consumes 2 ATP.
Seven rounds of /3-oxidation generate 8 acetyl CoA, 6 QH 2 (which yield 9 ATP via oxida-
tive phosphorylation), and 7 NADH (which yield 17.5 ATP). The fatty acid contains a
ds- /3,y double bond that is converted to a trans-a,f3 double bond, so the acyl-CoA
dehydrogenase- catalyzed reaction, which generates QH 2 , is bypassed in the fifth round.
Oxidation of the 8 acetyl CoA by the citric acid cycle yields 80 ATP. Therefore, the net
yield is 104.5 ATP equivalents.

4. When triacylglycerols are ingested in our diets, the hydrolysis of the dietary lipids occurs
mainly in the small intestine. Pancreatic lipase catalyzes the hydrolysis at the C-l and C-3
positions of triacylglycerol, producing free fatty acids and 2-monoacylglycerol. These mol-
ecules are transported in bile-salt micelles to the intestine, where they are absorbed by in-
testinal cells. Within these cells, the fatty acids are converted to fatty acyl CoA molecules,
which eventually form a triacylglycerol that is incorporated into chylomicrons for trans-
port to other tissues. If the pancreatic lipase is inhibited, the ingested dietary triglyceride
cannot be absorbed. The triglyceride will move through the digestive tract and will be ex-
creted without absorption.

734

SOLUTIONS Chapter 16

5. (a) Oleate has a cis- A 9 double bond, so oxidation requires enoyl-CoA isomerase (as in Step 2

of Figure 16.26).

(b) Arachidonate has cis double bonds at both odd (A 5 , A 11 ) and even (A 8 , A 14 ) carbons, so
oxidation requires both enoyl-CoA isomerase and 2,4-dienoyl-CoA reductase (as in Step 5
of Figure 16.26).

(c) This C 17 fatty acid contains a cis double bond at an even-numbered carbon (A 6 ), so oxi-
dation requires 2,4-dienoyl-CoA reductase. In addition, three enzymes are required to
convert the propionyl CoA product into succinyl CoA: propionyl-CoA carboxylase,
methylmalonyl-CoA racemase, and methylmalonyl-CoA mutase (Figure 16.25).

6. Even-chain fatty acids are degraded to acetyl CoA, which is not a gluconeogenic precursor.
Acetyl CoA cannot be converted directly to pyruvate because for every two carbons of acetyl
CoA that enter the citric acid cycle, two carbons in the form of two C0 2 molecules leave as
products. The last three carbons of odd-chain fatty acids, on the other hand, yield a molecule
of propionyl CoA upon degradation in the fatty acid oxidation cycle. Propionyl CoA can be
carboxylated and converted to succinyl CoA in three steps (Figure 16.25). Succinyl CoA can
be converted to oxaloacetate by citric acid cycle enzymes, and oxaloacetate can be a gluco-
neogenic precursor for glucose synthesis.

7. (a) The labeled carbon remains in H 14 COP; none is incorporated into palmitate. Although

H 14 COp is incorporated into malonyl CoA (Figure 16.2), the same carbon is lost as C0 2
during the ketoacyl-ACP synthase reaction in each turn of the cycle (Figure 16.5).

(b) All the even-numbered carbons are labeled. Except for the acetyl CoA that becomes C-15
and C-16 of palmitate, the acetyl CoA is converted to malonyl CoA and then to malonyl-
ACP before being incorporated into a growing fatty acid chain with the loss of C0 2 .

8. (a) Enoyl ACP reductase catalyzes the second reductive step in the fatty acid biosynthesis pathway,

converting a trans- 2,3 enoyl moiety into a saturated acyl chain, and uses NADPH as cofactor.

H O

I II

R — C = C — C — S — ACP

enoyl-A C pU NADPH + H@
reductase ^ ^

^NADP^

R — CH 2 — CH 2 — C — S — ACP

(b) Fatty acids are essential for membranes in bacteria. If fatty acid synthesis is inhibited,
there will be no new membranes and no growth of the bacteria.

(c) The fatty acid synthesis systems are different in animals and bacteria. Animals contain a
type I fatty acid synthesis system (FAS I) where the various enzymatic activities are local-
ized to individual domains in a large, multifunctional enzyme. In bacteria, each reaction
in fatty acid synthesis is catalyzed by a separate monofunctional enzyme. Understanding
some of the differences in these two systems, would allow for the design of specific inhibitors
of the bacterial FAS II.

9. Eating stimulates the production of acetyl CoA from the metabolism of carbohydrates (gly-
colysis and pyruvate dehydrogenase) and fats (FA oxidation). Normally, increased acetyl CoA
results in the elevation of malonyl CoA levels (acetyl CoA carboxylase reaction, Figure 16.2),
which may act to inhibit appetite. By blocking fatty acid synthase enzyme, C75 prevents the
removal of malonyl CoA for the synthesis of fatty acids, thereby elevating the levels of malonyl-
CoA and further suppressing appetite.

10 .

(a)

Carbohydrates

Glucose

Glycolysis

Pyruvate —

MITOCHONDRION
( ~ \
Citrate

Acetyl CoA

> Pyruvate

V J

Citrate

Acetyl CoA

Fatty acid
synthesis

Fatty acids

Chapter 16 SOLUTIONS 735

(b) The NADH generated by glycolysis can be transformed into NADPH by a variety of dif-
ferent reactions and pathways.

11. (a) Plentiful citrate and ATP levels promote fatty acid synthesis. High citrate levels activate

ACC by preferential binding and stabilization of the active dephosphorylated filamentous
form. On the other hand, high levels of fatty acyl Co As indicate that there is no further
need for more fatty acid synthesis. Palmitoyl CoA inactivates ACC by preferential binding
to the inactive protomeric dephosphorylated form.

(b) Glucagon and epinephrine inhibit fatty acid synthesis by inhibiting the activity of acetyl
CoA carboxylase. Both hormones bind to cell receptors and activate cAMP synthesis,
which in turn activates protein kinases. Phosphorylation of ACC by protein kinases con-
verts it to the inactive form, thus inhibiting fatty acid synthesis. On the other hand, the
active protein kinases catalyze phosphorylation and activation of triacylglycerol lipases
that catalyze hydrolysis of triacylglycerols, releasing fatty acids for (3 -oxidation.

12. (a) An inhibitor of acetyl-CoA acetylase will affect a key regulatory reaction for fatty acid

synthesis. The concentration of malonyl CoA, the product of the acetyl-CoA carboxylase-
catalyzed reaction, will be decreased in the presence of the inhibitor. The decrease in the
concentration of malonyl CoA will relieve the inhibition of carnitine acyltransferase I,
which is a key regulatory site for the oxidation of fatty acids. Thus, with an active carrier
system, fatty acids will be translocated to the mitochondrial matrix where the reactions of
/3 -oxidation occur. In the presence of an inhibitor of acetyl-CoA carboxylase, fatty acid
synthesis will decrease and (3 -oxidation will increase.

(b) CABI is a structural analog of biotin. Acetyl-CoA carboxylase is a biotin-dependent enzyme.

A biotin analog may bind in place of biotin and inhibit the activity of acetyl-CoA carboxylase.

13. The overall reaction for the synthesis of palmitate from acetyl CoA is the sum of two processes:

(1) the formation of seven malonyl CoA by the action of acetyl-CoA carboxylase and (2)
seven cycles of the fatty acid biosynthetic pathway.

7 Acetyl CoA + 7 C0 2 + 7 ATP > 7 Malonyl CoA + 7 ADP + 7 Pj

14. (a) Arachidonic acid is a precursor for synthesis of eicosanoids including “local regulators”
such as prostaglandins, thromboxanes, and leukotrienes (Figure 16.14). These regulators
are involved in mediation of pain, inflammation, and swelling responses resulting from
injured tissues.

(b) Both prostaglandins and leukotrienes are derived from arachidonate, which is released
from membrane phospholipids by the action of phospholipases. By inhibiting a phos-
pholipase, steroidal drugs block the biosynthesis of both prostaglandins and leukotrienes.
Aspirin-like drugs block the conversion of arachidonate to prostaglandin precursors by
inhibiting cyclooxygenase but do not affect leukotriene synthesis.

15. (a) O

o ch 2 — o— cr,

II I

r 2 — c — o— ch o

CH,— O— P — O — CH,

CHOH

CH 2 OH

(b) H H

O CH 2 — O — C = C — R,

-O — CH

ch 2 — o— p— ch,ch,nh,@

,©

I H

O CH — C = C — (CH 2 ) 12 — CH 3

II I H

R — C — NH — CH

CH 2 — o

736 SOLUTIONS Chapter 17

(COX-2)

16. Palmitate is converted to eight molecules of acetyl Co A labeled at C- 1 . Three acetyl CoA mol-
ecules are used to synthesize one molecule of mevalonate (Figure 16.17).

H 3 C — (CH 2 CH 2 ) 7 — COO°
Palmitate

8 H 3 C— C — S-CoA
Acetyl CoA

3 H 3 C—C — S-CoA
Acetyl CoA

Mevalonate

17. Both APHS and aspirin transfer an acetyl group to a serine residue on COX enzymes. Since
APHS is an irreversible inhibitor, it does not exhibit competitive inhibition kinetics even
though it acts at the active site of COX enzymes.

' O 1

(COX-2) — CH 2 0

^CH 2 C = C(CH 2 ) 3 CH 3

Irreversibly inhibited enzyme

Chapter 17 Amino Acid Metabolism

1. PSII contains the oxygen evolving complex and oxygen is produced during photosynthesis.
Since oxygen inhibits nitrogenase, the synthesis of 0 2 in hetocysts must be avoided. PSI is re-
tained because it can still generate a light-induced proton gradient by cyclic electron trans-
port and it is not involved in the production of 0 2 .

2. (a) Glutamate dehydrogenase + glutamine synthetase

NH 3 + Glutamate + ATP > Glutamine + ADP + Pj

(b) Glutamine synthetase + glutamate synthase

2 NH 3 + 2 Glutamate + 2 ATP > 2 Glutamine + 2 ADP + 2 Pj

The coupled reactions in (b) consume one more ATP molecule than the coupled reactions in (a).
Because the K m of glutamine synthetase for NH 3 is much lower than the K m of glutamate dehy-
drogenase for NH 4 ©, the coupled reactions in (b) predominate when NH 4 © levels are low. Thus,
more energy is spent to assimilate ammonia when its concentration is low.

3. The 15 N-labeled amino group is transferred from aspartate to a-ketoglutarate, producing
glutamate in a reaction catalyzed by aspartate transaminase (Figure 17.10). Since transami-
nases catalyze near- equilibrium reactions and many transaminases use glutamate as the
a- amino group donor, the labeled nitrogen is quickly distributed among the other amino
acids that are substrates of glutamate-dependent transaminases.

(a) a-Ketoglutarate + Amino acid ^

Oxaloacetate + Amino acid <
Pyruvate + Amino acid <

(b)

- Glutamate + a-Keto acid

- Aspartate + a-Keto acid
* Alanine + a-Keto acid

NAD(P)H, H

NAD(P)

-Ketoglutarate + NH 4

Glutamate

dehydrogenase

Chapter 17 SOLUTIONS 737

(Plants)

(Animals)

Serine — » O-acetylserine

(Sulf ide)S® >

Cysteine-SH (Fig 17.17)

Homoserine

Homocysteine-SH — » Methionine-S-CH 3

(Fig 17.11)

Methionine-S-CHo

Homocysteine-SH (Fig 17.35)

Serine

• Cystathionine (S)

Cysteine-SH (Fig 17.18)

6. (a) C-3 of serine is transferred to tetrahydrofolate during the synthesis of glycine, and C-2 is
transferred to tetrahydrofolate when glycine is cleaved to produce ammonia and bicarbonate.

,coo 0 — 0

0 I

H 3 N — 2 CH + Tetrahydrofolate <_

3 CH 2 OH

Serine

XOO

, © i

± h 3 n — 2 ch 2

+ 5,10-Methylenetetrahydrofolate + H 2 0

Glycine

COO

h 3 n— ch 2

+ Tetrahydrofolate + NAD® + H 2 0

5,10-Methylenetetrahydrofolate

Glycine

(b) Serine is synthesized from 3-phosphoglycerate (Figure 17.15), an intermediate of glycolysis.
C-3 of both 3-phosphoglycerate and serine is derived from either C-l or C-6 of glucose, and
C-2 of both 3-phosphoglycerate and serine is derived from either C-2 or C-5 of glucose.

7. (a)

COO

-CH

,©

/ \

h 3 c oh

(b)

®nh 3

ch 2 — ch-

coo

NADH + HC0 3 e

(c)

COO

0/™

h 2 i\t xh 2

\ /

h 2 c — ch 2

8. (a) Glutamic acid. PPI inhibits glutamine synthetase.

(b) Histidine biosynthesis pathway (Figure 17.23).

9. Aspartame is a dipeptide consisting of an asparate and a phenylalanine residue joined by a
peptide bond. This bond is eventually hydrolyzed inside the cell producing aspartate and
phenylalanine. Phenylketonuria patients must avoid any excess phenylalanine.

10. (a)

H,C

CH— CH 2

H,C

Leucine

-c-

-coo

H,C

Valine

-c-

-coo

h 3 c— h 2 c

Isoleucine

CH— CH— COO

H,C

(b) Lysine degradation pathway. a-Aminoadipate 8 - semialdehyde synthase is deficient
(Figure 17.39).

11 . (a) Alanine (c) Glycine

(b) Aspartate (d) Cysteine

12. The urea cycle does not operate in muscle, so ammonia from the deamination of amino acids
cannot be converted to urea. Because high concentrations of ammonia are toxic, ammonia is
converted to other products for disposal. In the first pathway, ammonia is incorporated into
glutamine by the action of glutamine synthetase (Figure 17.5). Glutamine can then be

NH 4 ® + H®

738

SOLUTIONS Chapter 18

transported to the liver or kidneys. The second pathway is the glucose-alanine cycle (Figure
17.45). Pyruvate accepts the amino group of amino acids by transamination, and the alanine
produced is transported to the liver where it can be deaminated back to pyruvate. The amino
group is used for urea synthesis, and the pyruvate can be converted to glucose.

13. Inhibition of nitric oxide synthase (NOS) can prevent excess amounts of nitric oxide from
being produced in cells lining the blood vessels. Nitric oxide causes relaxation of the vessels
and in excess amounts can cause reduced blood pressure leading to shock. Thiocitrulline and
S-methylthocitrulline inhibit NOS because they are unreactive analogs of the NOS reaction
product citrulline (Figure 17.25).

14. There are two reasons. Firstly, many of the amino acid biosynthesis pathways aren’t found in
humans, so there won’t be any metabolic diseases of nonexistent essential amino acid path-
ways. Secondly, the remaining pathways are probably crucial pathways during development so
that any defects in these pathways are likely to be lethal. This is the same reasoning that we used
to explain the lack of metabolic diseases in the sphingolipid biosynthesis pathways (Box 16.2).

15. The 21st, 22nd, and 23rd amino acids are N-formylmethionine, selenocysteine, and pyrrolysine.
N-formylmethionine and selenocysteine are synthesized during translation on aminoacylated
tRNA and not by the standard metabolic pathways covered in this chapter. Pyrrolysine may also
be synthesized on aminoacylated tRNA. The precursors are methionine, serine, and lysine.

16. The precursor in the serine biosynthesis pathway is 3-phosphoglycerate. This precursor can be de-
rived from glyceraldehyde-3-phosphate (G3P) in the glycolytic pathway, where the conversion is
associated with the gain of 1 ATP + 1 NADH. This gain must be subtracted from the total cost
of G3P synthesis. Therefore, the cost of making 3-phosphoglycerate is 24 — 3.5 = 20.5 ATP
equivalents, assuming that each NADH is equivalent to 2.5 ATPs. (The same cost can be derived
from the Calvin cycle pathway.) The serine biosynthesis pathway produces one NADH when
3-phosphoglycerate is oxidized to 3-phosphohydroxypyruvate, so the next cost of making serine
is 20.5 — 2.5 = 18 ATP equivalents. This value is identical to the value given in Box 17.3. (Note
that the transamination reaction in the serine biosynthesis pathway is cost-free.)

Alanine is made from pyruvate in a simple, cost-free, transamination reaction. The cost of
making pyruvate can be estimated from the conversion of 3-phosphoglycerate to pyruvate in
the glycolytic pathway. This conversion is associated with a gain of 1 ATP, so the cost of pyru-
vate is 20.5 — 1 = 19.5 ATP equivalents. Thus, the cost of synthesizing alanine is 19.5 ATP
equivalents, or 20 ATP equivalents when rounded to two significant figures. This value is the
same as that given in Box 17.3.

Chapter 18 Nucleotide Metabolism

1. (a)

NH,

TX>

Ribose

5-phosphate

(b)

Ribose

5-phosphate

(c)

Ribose

5-phosphate

See Figure 18.10 for the reactions in the pathway of UMP synthesis.

(d) Labeled C-2 from aspartate, which is incorporated into carbamoyl aspartate, appears at
C-6 of the uracil of UMP.

(e) The labeled carbon from HCO^, which is incorporated into carbamoyl phosphate, ap-
pears at C-2 of the pyrimidine ring of UMP.

(b) from r

HCOP HN^ XH

\ I II

6 CH
m • — V

(a) from C-2
of aspartate

Chapter 18 SOLUTIONS

739

2. Seven ATP equivalents are required. One ATP is cleaved to AMP when PRPP is synthesized
(Figure 18.3). The pyrophosphoryl group of PRPP is released in step 1 of the IMP biosynthetic
pathway and subsequently hydrolyzed to 2 Pi (Figure 18.5), accounting for the second ATP
equivalent. Five ATP molecules are consumed in steps 2, 4, 5, 6, and 7.

3. Purines: Reaction 3: GAR transformylase 1 0-formyl -THF, C-8 position.

Reaction 9: AICAR transformylase, 1 0-formyl -THF, C-2 position.
Pyrimidines: Thymidylate synthase, 5,1 0-methylene-THF, 5-CH 3 of thymidylate.

4. (a)

coo°

N — C — H
1

H 3 N — C — H

ch 2

o ch 2

ch 2

\ /

N = C

FUN — C v

Acivicin

Glutamine

(b) Acivicin inhibits glutamine-PRPP amidotransferase, the first enzyme in the purine
biosynthetic pathway, so PRPP accumulates.

(c) Acivicin inhibits the carbamoyl phosphate synthetase II activity of dihydroorotate syn-
thase that catalyzes the first step in the pyrimidine biosynthetic pathway.

5. (a) When /3-alanine is used instead of aspartate, no decarboxylation reaction (step 6 of the
E. coli pathway) would be required.

(b) 9

FIN CH

i Ji

Ribose

5-phosphate

(b) Synthesis of DNA requires certain ratios of A, T, G and C. If dTTP levels are higher than
necessary, dTTP will act to decrease its own synthesis pathway by inhibiting the conver-
sion of dCMP to dUMP by dCMP deaminase. dUMP is the precursor to dTMP (thymidy-
late synthase, Figure 18.16), and the subsequent conversion to dTDP and dTTP (needed
for DNA synthesis). On the other hand, if dCTP levels are high, activation of dCMP
deaminase will lead to an increased conversion of dCMP to dUMP and this diverts any
dCMP that might have been converted to more dCTP by phosphorylation (Figure 18.20).

7. Four ATP equivalents are required. One ATP equivalent is required for the synthesis of PRPP
from ribose 5-phosphate (Figure 18.3). Carbamoyl phosphate synthesis requires 2 ATP (Figure
18.10, step 1). One ATP equivalent is consumed in step 5, when PP^ is hydrolyzed to 2 P*.

8. In the absence of adenosine deaminase, adenosine and deoxyadenosine are not degraded via in-
osine and hypoxanthine to uric acid (Figure 18.19 and 18.21). This leads to an increase in the
concentration of deoxyadenosine, which can be converted to dATP. High concentrations of
dATP inhibit ribonucleotide reductase (Table 18.1). The inhibition of ribonucleotide reductase
results in decreased production of all deoxynucleotides and therefore inhibits DNA synthesis.

9. Glutamine-PRPP amidotransferase is the first enzyme and the principal site of regulation in the de
novo pathway to IMP (Figure 18.5). In humans, PRPP is both a substrate and a positive effector of
this enzyme. An increase in the cellular levels of PRPP due to increased PRPP synthetase activity
will therefore enhance the activity of the amidotransferase. This will result in an increased synthe-
sis of IMP and other purine nucleosides and nucleotides. Overproduction of purine nucleotides
and subsequent degradation can lead to elevated uric acid levels characteristic of gout.

10. (a) ATP (b) ATP (c) ATP (d) GTP (e) UTP (f) GTP (g) C TP (h) UTP (i) ATP (j) IMP (k) IMP

11. Purines and pyrimidines are not significant sources of energy. The carbon atoms of fatty
acids and carbohydrates can be oxidized to yield ATP, but there are no comparable energy-
yielding pathways for nitrogen-containing purines and pyrimidines. However, the NADH
produced when hypoxanthine is converted to uric acid may indirectly generate ATP via

740

SOLUTIONS Chapter 19

oxidative phosphorylation. The degradation of uracil and thymine yields acetyl CoA and suc-
cinyl CoA, respectively, which can be metabolized via the citric acid cycle to generate ATP.

12. The sugar D-ribose exists as an equilibrium mixture of u-D-ribopyranose, cr-D-ribofuranose,
/3-D-ribopyranose, and /3-D-ribofuranose. These forms freely interconvert with each through
the open-chain form (Section 8.2).

13. Xanthine is 2,6-dioxopurine; hypoxanthine is 6-oxopurine; orotate is 2,4-dioxo-6-carboxyl-
pyrimidine.

14. SAICAR synthetase + adenylosuccinate lyase in the IMP biosynthesis pathway (Figure 18.5)
and argininosuccinate synthetase + argininosuccinate lyase in the arginine biosynthesis
pathway (urea cycle: Figure 17.43).

Chapter 19 Nucleic Acids

1. In the a helix, hydrogen bonds form between the carbonyl oxygen of one residue and the
amine hydrogen four residues, or one turn, away. These hydrogen bonds between atoms in
the backbone are roughly parallel to the axis of the helix. The amino acid side chains, which
point away from the backbone, do not participate in intrahelical hydrogen bonding. In double-
stranded DNA, the sugar-phosphate backbone is not involved in hydrogen bonding. Instead,
two or three hydrogen bonds, which are roughly perpendicular to the helix axis, form between
complementary bases in opposite strands.

In the a helix, the individual hydrogen bonds are weak, but the cumulative forces of these
bonds stabilize the helical structure, especially within the hydrophobic interior of a protein
where water does not compete for hydrogen bonding. In DNA, the principal role of hydrogen
bonding is to allow each strand to act as a template for the other. Although the hydrogen
bonds between complementary bases help stabilize the helix, stacking interactions between
base pairs in the hydrophobic interior make a greater contribution to helix stability.

2. If 58% of the residues are (G + C), 42% of the residues must be (A + T). Since every A pairs
with a T on the opposite strand, the number of adenine residues equals the number of
thymine residues. Therefore, 21%, or 420, of the residues are thymine (2000 X 0.21 = 420).

3. (a) The base compositions of complementary strands of DNA are usually quite different. For

example, if one strand is poly dA (100% A), the other strand must be poly dT (100% T).
However, since the two strands are complementary, the amount of (A + T) must be the
same for each strand, and the amount of (G + C) must be the same for each strand.

(b) (A + G) = (T + C). Complementarity dictates that for every purine (A or G) on one
strand, there must be a pyrimidine (T or C) on the complementary strand.

4. Since the DNA strands are anti- parallel, the complementary strand runs in the opposite di-
rection. The sequence of the double-stranded DNA is

AT CGCGTAACAT GGATT CGG
TAGCGCATTGTACCTAAGCC

By convention, DNA sequences are written in the 5' — » 3' direction. Therefore, the sequence
of the complementary strand is

CCGAAT CCAT GTTACGCGAT

5. The stability of the single- stranded helix is largely due to stacking interactions between adja-
cent purines. Hydrophobic effects also contribute, since the stacked bases form an environ-
ment that is partially shielded from water molecules.

7. There will be two discrete melting points separated by a plateau. When the extra strand of
poly dT is released, the absorbance of the solution at 260 nm will increase as the stacked bases
leave the largely hydrophobic interior of the triple helix. A second increase in the absorbance
occurs when the remaining two DNA strands denature.

Chapter 19 SOLUTIONS

741

8 .

10 .

11 .

12 .

13.

14.

15.

16.

17.

18 .

The sequence is

5' ACG CACGUAUA UGUACU UAUACGUGG CU 3'

The underlined sequences are palindromic.

The main products will be a mixture of mononucleotides and pieces of single- stranded DNA
approximately 500 bp in length. A piece of DNA with an enzyme molecule bound at each end
will be degraded until the two strands can no longer base-pair; at that point the single strands
cease to be a substrate for the enzyme.

In the 30 nm fiber, DNA is packaged in nucleosomes, each containing about 200 bp of DNA;
therefore, the DNA in a nucleosome has a molecular weight of 130,000 (200 X 650 = 130,000).
Assuming there is one molecule of histone HI per nucleosome, the molecular weight of the pro-
tein component of the nucleosome would be 129,800.

Histone HI

21,000

Histone H2A (X2)

28,000

Histone H2 B (X2)

27,600

Histone H3 (X2)

30,600

Histone H4 (X2)

22.600

Total

129,800

Thus, the ratio by weight of protein to DNA is 129,800:130,000, or approximately 1:1.

Nucleosomes are composed of histones plus 200 base pairs of DNA. Since you inherited half
your chromosomes from your mother, the oocyte contained

(3.2 X 10 9 bp) x

1 nucleosome
200 bp

= 8 X 1 0 6 nucleosomes

(You inherited no nucleosomes from your father since nucleosomes are replaced by small,
positively charged polypeptides during spermatogenesis.)

(a) pdApdGpdT + pdC

(b) pdAp + dGpdTpdC

Since the supercoiled plasmid DNA is in equilibrium with relaxed DNA containing short un-
wound regions, the Aspergillus enzyme will slowly convert the DNA into nicked circles. Even-
tually the enzyme will convert the relaxed circles into unit-length linear fragments of
double-stranded DNA.

Yes. The sugar-phosphate backbone in both RNA and DNA contains phosphodiester bonds
that link the sugar residues.

pppApCpUpCpApUpApGp + CpUpApUpGp + ApGp + U

Bacteriophages have evolved several mechanisms to protect their DNA from restriction en-
donucleases. In general, bacteriophage DNA contains few restriction sites. Restriction en-
donuclease recognition sites are strongly selected and any mutations that alter these sites will
be favored. In addition, restriction sites are often methylated, as in the bacterial chromosome.
This is presumably due to a fortuitous event in the distant past when the phage DNA became
methylated before it could be cleaved.

Some bacteriophages incorporate modified nucleotides into their DNA. The modified nu-
cleotides (e.g., 5-hydroxymethylcytosine in bacteriophage T4) are not recognized by restric-
tion endonucleases.

Phage genomes may also encode an enzyme that inactivates restriction endonucleases, or
they may encode proteins that bind to restriction sites to prevent cleavage.

(a) The probability can be estimated from the probability of each nucleotide in the Hind III
restriction site. (G = C = 0.18 and A = T = 0.32)

For the sequence AAGCTT there will be, on average, one Hindlll site every
1/(0. 32) (0.32)(0. 18) (0.18) (0.32) (0.32) = 2943 bp
Thus, in a 100 Mb genome there will be, on average,

100,000/2943 = 33,070 sites

(b) 24,414

Although the recognition sites for Bglll and BamHl differ, the enzymes produce fragments
with identical sticky ends. These fragments can be ligated as easily as fragments produced by
a single enzyme.

742

SOLUTIONS Chapter 20

Bgl 1 1 'vw' A G A T C T 'vw

'vw' T C T A G A

BamYW 'wv'G G ATCC^

vw C C T A G G ' vx/x/ '

19. Restriction enzymes present in normal host cells might cleave newly introduced recombinant
molecules, making it impossible to clone certain fragments of DNA. Using a host strain that
does not make restriction endonucleases avoids this problem.

A mutation in RecA reduces recombination, thereby preventing the rearrangement of recom-
binant DNA molecules during propagation in the host cells. Rearrangement is often a problem,
particularly when the cloned fragment of DNA contains repetitive sequences that can serve
as sites for homologous recombination.

Chapter 20 DNA Replication, Repair, and Recombination

1. (a) Two replication forks form at the origin of replication and move in opposite directions
until they meet at a point opposite the origin. Therefore, each replisome replicates half the
genome (2.6 X 10 2 3 4 * 6 base pairs). The time required to replicate the entire chromosome is

2.6 X 1 0 6 base pairs
1 000 base pairs s _1

= 2600 s = 43 min and 20 s

(b) Although there is only one origin (O), replication can be reinitiated before the previous
replication forks have reached the termination site. Thus, the chromosome can contain
more than two replication forks. Replication of a single chromosome still requires ap-
proximately 43 minutes, but completed copies of each chromosome can appear at
shorter intervals, depending on the rate of initiation.

Replication forks initiated
before completion of first
round of DNA replication

2. T4 DNA polymerase should be an early gene product because it is required for replication of
the viral genome.

3. (a) The single-stranded DNA template used for DNA synthesis in vitro can form secondary

structures such as hairpins. SSB prevents the formation of double-stranded structure by
binding to the single- stranded template. SSB thus renders the DNA a better substrate for
DNA polymerase.

(b) The yield of DNA in vitro is improved at higher temperatures because formation of sec-
ondary structure in the template is less likely. A temperature of 65°C is high enough to
prevent formation of secondary structure but not high enough to denature the newly
synthesized DNA. DNA polymerases from bacteria that grow at high temperatures are
used because they are active at 65°C, a temperature at which DNA polymerases from
other bacteria would be inactive.

4. Extremely accurate DNA replication requires a proofreading mechanism to remove errors in-

troduced during the polymerization reaction. Synthesis of an RNA primer by a primase,

which does not have proofreading activity, is more error prone than DNA synthesis. However,

Chapter 20 SOLUTIONS 743

because the primer is RNA, it can be removed by the 5' — > 3' exonuclease activity of DNA
polymerase I and replaced with accurately synthesized DNA when Okazaki fragments are
joined. If the primer were composed of DNA made by a primase without proofreading activ-
ity, it would not be removed by DNA polymerase I and the error rate of DNA replication
would be higher at sites of primer synthesis.

5. (a) In the hypothetical nucleotidyl group transfer reaction, the nucleophilic 3 '-hydroxyl
group of the incoming nucleotide would attack the triphosphate group of the growing
chain. Pyrophosphate would be released when a new phosphodiester linkage was formed.

©r

\©

©r

\©

Incoming nucleoside
triphosphate

Growing chain

3' DNA

(b) If the hypothetical enzyme had 5' — > 3' proofreading activity, removal of a mismatched
nucleotide would leave a 5 '-monophosphate group at the end of the growing chain. Fur-
ther DNA synthesis, which would require a terminal triphosphate group, could not occur.

6. Topisomerase II or gyrase relieves supercoiling ahead of and behind the replication fork. If
this enzyme is inhibited, the unwinding of the parental DNA cannot occur. Therefore, the
DNA of the E. coli cannot be replicated.

7. (a) Assume that the genome is one large linear molecule of DNA and that the origin of repli-
cation is at the midpoint of this chromosome. Since the replication forks move in oppo-
site directions, 60 base pairs can be replicated per second. The time required to replicate
the entire genome would be

1 .65 x 1 0 8 base
60 base pairs s

pairs

= 2.75 X 1 0 6 s = 764 h = 32 days

744

SOLUTIONS Chapter 20

(b) Assuming that the 2000 bidirectional origins are equally spaced along the DNA molecule
and that initiation occurs simultaneously at all origins, the rate would be
2000 X 2 X 30 base pairs per second, or 1.2 X 10 5 base pairs per second. The time re-
quired to replicate the entire genome would be

1 .65 x 10 8 base pairs
1.2 x 10 5 base pairs s -1

1 375 s = 23 min

(c) Assume that the origins are equally spaced and that initiation at all origins is simultane-
ous. The required rate of replication is

1 .65 x 1 0 8 base pairs
300 s

= 5.5 x 10 5 base pairs s 1

Bidirectional replication from each fork proceeds at an overall rate of 60 base pairs per sec-
ond. The minimum number of origins would be

5.5 X 10 5 base pairs s 1
60 base pairs s -1 origin -1

91 70 origins

8. The modified G can no longer form a productive Watson-Crick base pair with C but can now
base-pair with T. Therefore, one of the daughter strands of DNA will contain a T across from
the modified base. After further rounds of replication, the T will base-pair with A and what
was originally a G/C base pair will have mutated into an A/T base pair.

9. Ultraviolet light can damage DNA by causing dimerization of thymidylate residues. One
mechanism for repairing thymine dimers is enzymatic photoreactivation, catalyzed by DNA
photolyase. This enzyme uses energy from visible light to cleave the dimer and repair the
DNA. Thus, cells that are exposed to visible light following ultraviolet irradiation are better
able to repair DNA than cells kept in the dark.

10 . (a) DNA from a dut~ strain will appear normal because the Ung enzyme will remove any

uracil that gets incorporated.

(b) DNA from a dut ~ , ung~ strain will contain dU residues in the place of some dT residues.

11. The DNA repair enzyme uracil N-glycosylase removes uracil formed by the hydrolytic deam-
ination of cytosine. Because the enzyme does not recognize thymine or the other three bases
normally found in DNA, it cannot repair the damage when 5-methylcytosine is deaminated
to thymine.

12 . High mutation rates occur at methylcytosine- containing regions because the product of
deamination of 5-methylcytosine is thymine, which cannot be recognized as abnormal.
When the mismatched T/G base pair that results from deamination of methylcytosine is re-
paired, the repair enzymes may delete either the incorrect thymine or the correct guanine.
When the guanine is replaced by adenine, the resulting A/T base pair is a mutation.

Normal

sequence

Parental

strand

Mutation

Daughter

strand

Chapter 21 SOLUTIONS

745

13. Proofreading during replication results in excision of 99% of misincorporated nucleotides,
thus reducing the overall error rate to 10 -7 . Of those errors that escape the proofreading step,
a further 99% are corrected by repair enzymes. The overall mutation rate is therefore 10 -9 .

14. Yes. The E. coli enzyme DNA ligase is required to seal the nicks left in the DNA strands follow-
ing DNA repair. This enzyme has a strict requirement for NAD®.

15. The dimers can be removed by excision repair. UvrABC endonuclease removes a 12-13
residue segment containing the pyrimidine dimer. The DNA oligonucleotide is removed with
the help of a helicase. The gap is filled by the action of DNA polymerase I, and the nick sealed
by the action of DNA ligase. The dimers can also be repaired through direct repair. DNA pho-
tolyase binds to the distorted double helix at the site of the dimer. As the DNA-enzyme com-
plex absorbs light, the dimerization reaction is reversed.

16. The repair enzymes need an undamaged template in order to repair mutations in DNA. If both
strands of the DNA molecule have been damaged, there is not a template to use for repair.

17. The proteins that catalyze strand exchange recognize regions of high sequence similarity and
promote formation of a triple-stranded intermediate in which the invading strand base-pairs
with a complementary strand. This pairing would not be possible if the sequences of the two
DNA molecules were different.

18. DNA polymerase III is a component of the replisome that synthesizes the leading strand and
the lagging strand during replication of the E. coli chromosome. DNA polymerase I is re-
quired to remove the short RNA primers on the lagging strand.

Chapter 21 Transcription and RNA Processing

1. (a) Since the rate of transcription is 70 nucleotides per second and each transcription com-
plex covers 70 base pairs of DNA, an RNA polymerase completes a transcript and leaves
the DNA template each second (assuming that the complexes are densely packed). There-
fore, when the gene is loaded with transcription complexes, 60 molecules of RNA are pro-
duced per minute.

(b) Since each transcription complex covers 70 base pairs, the maximum number of
complexes is

6000 base pairs

— — : : : — : : — = 86 transcription complexes

70 base pairs transcription complex

2. (a) Since the average E. coli gene is 1 kb (1000 bp) long, 4000 genes account for 4000 kb of
DNA. The percentage of DNA that is not transcribed is

500 kb
4600 kb

X 100% = 10.9%

Most of the nontranscribed DNA consists of promoters and regions that regulate tran-
scription initiation.

(b) Since the gene products in mammals and bacteria are similar in size, the amount of DNA
in the exons of a typical mammalian gene must also be 1000 bp. The total amount of
DNA in exons is

5 X 1 0 4 genes X 1 .0 kb gene 1 = 5 X 1 0 4 kb

This DNA represents about 1.7% of the mammalian genome.

5 x 1 0 4 kb
3 x 1 0 6 kb

x 100% = 1.7%

The remaining 97.5% of DNA consists of introns and other sequences.

3. No. It is extremely unlikely that the eukaryotic genes promoter will contain the correct sequences
in the correct location to permit accurate initiation by the prokaryotic RNA polymerases.
Likewise, it is extremely unlikely that the prokaryotic gene s promoter will contain the correct se-
quence in the correct location to permit accurate initiation by RNA polymerase II.

4. No. A typical eukaryotic triose phosphate isomerase gene contains introns. The prokaryotic
cell contains no spliceosomes and therefore will not be able to correctly process the primary
transcript. Therefore, translation of the RNA will yield an aberrant protein fragment.

5. (a) In the presence of both lactose and glucose, the lac operon is transcribed at a low level be-
cause lac repressor forms a complex with allolactose (an isomer of lactose). Because the

746

SOLUTIONS Chapter 21

allolactose-repressor complex cannot bind to the promoter region of the lac operon, the
repressor does not prevent initiation of transcription.

(b) In the absence of lactose, no allolactose is formed. Thus, lac repressor binds near the lac
operon promoter and prevents transcription.

(c) When lactose is the sole carbon source, the lac operon is transcribed at the maximum
rate. In the presence of allolactose, transcription is allowed since lac repressor does not
bind to the promoter region of the lac operon. Also, in the absence of glucose, the tran-
scription rate increases because cAMP production increases, making more CRP-cAMP
available to bind to the promoter region of the lac operon. The absence of the repressor
and the enhancement of transcription initiation by CRP-cAMP allow the cell to synthe-
size the quantities of enzymes required to support growth when lactose is the only car-
bon source.

6. Since the wild-type lac promoter is relatively weak, maximal transcription requires the activa-
tor CRP. The UV5 mutations alters the —10 region such that it now resembles the consensus
— 10 sequence, making it a much stronger promoter. In the absence of the lac repressor, the
promoter is independent of CRP.

7. 32 P appears only at the 5' end of mRNA molecules that have ATP as the first residue. It does
not appear in any other residues because pyrophosphate, which includes the (3 -phosphoryl
group, is released when nucleoside triphosphates are added to the 3 ' end of a growing RNA
chain (Figure 21.3).

When the 5' end of mRNA is capped, only the y-phosphoryl group of the initial residue is
removed when the cap forms. The (3 -phosphoryl group, which contains the label, is retained
and receives the GMP group from GTP (Figure 21.26).

8. The lack of proofreading activity in RNA polymerase makes the error rate of transcription
greater than the error rate of DNA replication. However, the defective RNA molecules pro-
duced are not likely to affect cell viability because most copies of RNA synthesized from a
given gene are normal. In the case of defective mRNA, the number of defective proteins is
only a small percentage of the total number of proteins synthesized. Also, mistakes made dur-
ing transcription are quickly eliminated since most mRNA molecules have a short half-life.

9. During maturation, eukaryotic mRNA precursors are modified at their 3' ends by the addi-
tion of a poly A tail. When a mixture of components from a cell extract is passed over the col-
umn, the poly A tail will hybridize with oligo dT on the column. The other components in the
cell extract will pass through the column. The bound mature mRNA with the poly A tail is re-
moved from the column by changing the pH or the ionic strength of the buffer. This will dis-
rupt the hydrogen bonds between the A and T nucleotides.

10. (a) A much lower concentration of rifampicin stopped the growth of the wild-type E. coli

(<5 fig/ mL) as compared to the concentration of rifampicin that stopped the growth of
the mutant (>50 fig/mL).

(b) RNA polymerase consists of a core enzyme with a stoichiometry of a 2 /3l3 , (x) that partici-
pates in many of the transcription reactions. The large (3 and [3 ' subunits make up the
active site of the enzyme.

(c) The rifampicin-resistant bacteria could arise from mutations that occur in the gene for
the /3 subunit of RNA polymerase.

11. Since either strand can serve as a template, two mRNA molecules can be transcribed from this
DNA segment. When the bottom strand is the template, the mRNA sequence is complemen-
tary to the bottom strand.

5' 'VW' £
3 r 'vwr> Q

r GCTAAGATCTGACTa
c b a g

G C T C

L C G A T T CT AGACT G A

r G C UAAGAUCUG A
mRNA 5' C y

Direction of transcription '

C 'vw' 3 r

Q 'VW'

5'-wv.cCGGCUAAGAUCUGACUAGC-wv3'

mRNA

Chapter 22 SOLUTIONS

747

When the top strand is the template, the mRNA sequence is complementary to the top
strand.

HO,

Direction of transcription

"AU U C UAGAC U G A 1

5' mRNA

5 r / WV' £

3 r G

G GCTAAGATCTGACT a

c cgattctagactga t

£ 'VW'

Q 'va / vr

5 , 'WV'GCUAGUCAGAUCUUAGCCGG'W^3 /

mRNA

12. A gene was defined as a DNA sequence that is transcribed. By this definition, the entire ribo-
somal RNA operon is a gene. However, it is sometimes more convenient to restrict the term
gene to the segment of RNA that encodes a functional product, for example, one of the enzymes
encoded by the lac operon. The operon in Figure 21.25 therefore contains tRNA and 16S, 23S,
and 5S rRNA genes. The DNA sequences between these genes, although transcribed, are not
considered part of any gene.

13. The genomic DNA sequence provides an accurate rendition of the primary RNA sequence as
expected. However, sequencing a purified tRNA reveals that many of the nucleotides have
been specifically modified post- transcriptionally. The same is true for eukaryotes.

14. The gene for triose phosphate isomerase in maize contains about 3400 base pairs. If the
spliceosome assembles at the first intron, then 2900 base pairs remain to be transcribed. The
time required to transcribe 2900 base pairs is 97 seconds (2900 nucleotides 30 nucleotides
per second) . If the spliceosome assembles immediately after transcription of the first intron,
and if splicing cannot begin until transcription of the entire gene is complete, the spliceo-
some must be stable for at least 97 seconds.

15. The CRP-cAMP binding site probably overlaps the promoter of the gene. When CRP-cAMP
binds, the promoter is blocked and transcription cannot occur.

16. When the sequence of the 5' or 3' splice site or the branch point is altered by mutation, proper
splicing cannot occur and no functional mRNA can be produced.

17. Yes. Once the U2 snRNP binds to the branch site it will occlude the U5 snRNP from binding to
the 3' splice acceptor and interfere with splicing. Furthermore, the deletion will have removed
a large part of the pyrimidine stretch required for binding to the 3' splice site. Both of these
will prevent proper mRNA processing and the aberrant RNA will not be properly translated.

Chapter 22 Protein Synthesis

1 . One strand of DNA has three different overlapping reading frames, therefore a double- stranded
DNA has six reading frames. This can be seen by examining the DNA sequence beginning at the
5' end of each strand and marking off the triplet codons. This identifies one reading frame on
each strand. Now start at the second nucleotide in from the 5' ends and mark off triplet codons;
that is reading frame 2. The third reading frame on each strand begins at the third nucleotide in
from the 5 ' ends. The “fourth” reading frame is identical to the first — test this for yourself.

Using similar logic, it follows that if the genetic code were read in codons four nucleotides
in length, then one strand of DNA could be read in four different reading frames and
therefore a double-stranded piece of DNA would contain eight reading frames (four on
each strand).

2. Each mRNA sequence could be translated in three different reading frames. For the first
mRNA sequence, the possible codons and polypeptide sequences are

Reading Frame 1 5'

'w^CCGGCUAAGAUCUGACUAGC'wxa 3 /
— Pro — Ala — Lys — lie stop

Reading Frame 2 5 'waCCGGCUAAGAUCUGACUAGC^ 3'
— Ala — Leu — Arg — Ser — Asp stop

Reading Frame 3 5'

^CCGGCUAAGAUCUGACUAGC^3'
— Gly STOP

748

SOLUTIONS Chapter 22

For the second mRNA sequence, the possible codons and polypeptide sequences are

Reading Frame 1
Reading Frame 2
Reading Frame 3

5 , 'w^GCUAGUCAGAUCUUAGCCGG'w^3 /
— Ala — Ser — Gin — lie — Leu — Ala — Gly —

5' vw>g CLLA (TuC A~GA U~CU UA~G cTc~G 3'
— Leu — Val — Arg — Ser stop

5'^gc UAG UCA GA~U CLMJ AcTc CGG ^ 3'
STOP

Since only a reading frame without a stop codon can encode a polypeptide, the second
mRNA sequence corresponds to the actual transcript. The sequence of the encoded polypep-
tide is -Ala-Ser-Gln-Ile-Leu-Ala-Gly-

3. Two phosphoanhydride bonds are hydrolyzed for each amino acid activated by an aminoacyl-
tRNA synthetase.

Amino acid + tRNA + ATP > Aminoacyl - tRNA + AMP + PPj

PPi + H 2 0 > 2 Pi

The rest of the energy needed to synthesize the protein is provided by hydrolysis of GTP: one
“high energy” bond is hydrolyzed in the formation of the 70S initiation complex, another
during the insertion of each aminoacyl-tRNA into the A site of the ribosome, and another at
each translocation step. Since the initial methionyl-tRNA is inserted into the P site, 599 new
insertions and 599 translocations occur during the synthesis of a 600-residue protein. Finally,
one phosphoanhydride bond is hydrolyzed during release of the completed polypeptide
chain from the ribosome. The total number of phosphoanhydride bonds hydrolyzed during
synthesis of the protein is

Activation (600 X 2)

1200

Initiation

Insertion

599

Translocation

599

Termination

Total

2400

4. The answer depends on your frame of reference. For example, relative to the ribosome, the
mRNA and both tRNAs get translocated by one triplet codon. Relative to the mRNA, it is the
ribosome that is shifted by three nucleotides.

5. The region of the mRNA molecule upstream of the true initiation codon contains the purine-
rich Shine-Dalgarno sequence, which is complementary to a pyrimidine- rich sequence at the
3' end of the 16S rRNA component of the 30S ribosomal subunit (Figure 22.17). By cor-
rectly positioning the 30S subunit on the mRNA transcript, the Shine-Dalgarno sequence
allows fMct-tRNAf 1 ^ to bind to the initiation codon. Once protein synthesis begins, all sub-
sequent methionine codons are recognized by Met-tRNA Met .

6. No, because proper translation initiation in an E. coli cell requires a Shine-Dalgarno sequence
located in the 5 ' untranslated region of the mRNA. Since eukaryotic ribosomes do not have
this requirement, it is extremely unlikely that an mRNA from a plant would fortuitously con-
tain a Shine-Dalgarno sequence in the proper location.

If, however, the part of the gene encoding the plant mRNA were fused to a bacterial Shine-
Dalgarno sequence, then the open reading frame for the plant protein would be properly
translated in the bacterial cell.

7. The transcript of each rRNA gene is an rRNA molecule that is directly incorporated into a ri-
bosome. Thus, multiple copies of rRNA genes are needed to assemble the large number of
ribosomes that the cell requires. In contrast, the transcript of each ribosomal protein gene is
an mRNA that can be translated many times. Because of this amplification of RNA to protein,
fewer genes are needed for each ribosomal protein than for rRNA.

8. Possible suppressor tRNA species include all those that recognize codons differing from UAG
by a single nucleotide, that is, tRNAs whose anticodons differ by a single nucleotide from the
sequence CUA, which is complementary to the stop codon UAG. tRNA Gln , tRNA Lys , and
tRNA Gln all recognize codons that differ only at the first position (codons CAG, AAG, and
GAG, respectively). tRNA Leu , tRna Ser , and tRNA Trp recognize codons that differ only at the
second position (UUG, UCG, and UGG, respectively). tRNA Tyr recognizes codons that differ
only at the third position (UAU or UAC).

Chapter 22 SOLUTIONS

749

A cell that contains a suppressor tRNA can survive despite the loss of a normal tRNA because
the cell also contains isoacceptor tRNA molecules that carry the same amino acid. Although
the suppressor tRNA may occasionally insert an amino acid at a normally occurring stop
codon, the resulting protein, which is larger than the normal gene product, is usually not
lethal to the cell. In fact, strains of E. coli that contain suppressor tRNAs do survive but are
often not as healthy as wild-type strains.

9. (a) Aminoacyl-tRNA synthetases — the enzymes that bind to tRNAs and catalyze aminoacy-

lation.

(b) IF-2 in bacteria and eIF-2 in eukaryotes, a protein that binds to aminoacylated initiator-
tRNA and loads it into the ribosome’s P site during translation initiation.

(c) EF-Tu in bacteria and EF-la in eukaryotes — a protein that binds to charged tRNAs and
loads them into the ribosome’s A site during polypeptide elongation.

(d) Ribosomes. These large complexes of RNA and protein contain two sites that can bind
specifically to tRNAs, the A site and the P site.

(e) mRNA — tRNAs bind to mRNA through codon- anticodon hydrogen bonds.

The enzymes that modify specific residues on individual tRNAs during the maturation
process must also be able to bind to tRNAs.

10. Under normal circumstances, when the translation machinery encounters UGA in RF-2
mRNA, RF-2 recognizes the stop codon and terminates protein synthesis. When the cellular
concentration of RF-2 is low, however, the ribosome pauses at the termination codon, shifts
frame, and continues translating the RF-2 mRNA to produce the full-length functional pro-
tein. Thus, the presence of the stop codon encourages translational frameshifting in the ab-
sence of RF-2 and allows RF-2 to regulate its own production.

11. (a) If the entire leader region were deleted, attenuation would not be possible, and transcrip-

tion would be controlled exclusively by trp repressor. The overall rate of transcription of
the trp operon would increase.

(b) If the region encoding the leader peptide were deleted, transcription would be controlled
exclusively by trp repressor. Deletion of the sequence encoding the leader peptide would
remove Sequence 1, thus allowing the stable 2-3 hairpin to form. Since neither the pause
site (1-2 hairpin) nor the terminator (3-4 hairpin) could form, initiated transcripts
would always continue into the trp operon.

(c) If the leader region did not contain an AUG codon, the operon would be rarely tran-
scribed. Because of the absence of the initiation codon, the leader peptide would not be
synthesized, and 1-2 hairpins and 3-4 hairpins would almost always form, leading to ter-
mination of transcription.

12. No, this is difficult to imagine. One of the important features of the attenuation model is that
one or more codons in the leader peptide usually encode the amino acid that is synthesized by
that operon. It is the relative shortage or abundance of particular aminoacylated tRNAs that
modulates the attenuation. The products of the lac operon are not directly involved in amino
acid biosynthesis, so we would not expect cellular levels of one class of aminoacylated tRNAs
to vary with the activity of the operon.

13. The presence of codons specifying valine and leucine in the leader regions of isoleucine oper-
ons suggests that a scarcity of these amino acids would promote transcription of the genes for
isoleucine biosynthesis. Many of the enzymes required to synthesize isoleucine are also re-
quired in the pathways to valine and leucine (Section 18.5A). Thus, even when the isoleucine
concentration is high, a low concentration of valine or leucine ensures that transcription of
the isoleucine operon does not terminate prematurely.

14. As the newly synthesized protein is extruded from the ribosome, the N-terminal signal
peptide is recognized and bound by a signal-recognition particle (SRP). Further transla-
tion is inhibited until the SRP binds to its receptor on the cytosolic face of the endoplas-
mic reticulum. Ribophorins anchor the ribosome to the endoplasmic reticulum. When
translation resumes, the polypeptide chain passes through a pore into the lumen. If the
polypeptide does not pass completely through the membrane, the result is an integral
membrane protein with its N-terminus in the lumen of the endoplasmic reticulum and its
C-terminus in the cytosol.

Glycosylation of specific residues takes places in the lumen of the endoplasmic reticulum and
in the Golgi apparatus. The protein, still embedded in the membrane, is transported between
the endoplasmic reticulum and the Golgi apparatus in transfer vesicles that bud off the endo-
plasmic reticulum.

750

SOLUTIONS Chapter 22

Secretory vesicles transport the fully glycosylated protein from the Golgi apparatus to the
plasma membrane. When the vesicles fuse with the plasma membrane, the N - terminal por-
tion of the protein, which was in the lumen, is now exposed to the extracellular space, and the
C-terminal portion remains in the cytosol.

15. Yes. A hydrophobic secretion signal sequence located at the AT- terminus of a protein is neces-
sary and sufficient for entry into the cells secretory pathway.

16. The initiator tRNA anticodon pairs with GUG by forming a G/U base pair between the 5'
nucleotide of the codon and the 3' position of the anticodon.

3' 5'

Initiator tRNA anticodon y A c

mRNA codon 5' G U G 3'

This interaction is unrelated to wobble since the 5' position of the anticodon is the wobble
position.

Glossary of Biochemical Terms

A site. Aminoacyl site. The site on a ribo-
some that is occupied during protein synthe-
sis by an aminoacyl-tRNA molecule,
acceptor stem. The sequence at the 5' end
and the sequence near the 3' end of a tRNA
molecule that are base paired, forming a stem.
The acceptor stem is the site of amino acid at-
tachment. Also known as the amino acid stem,
accessory pigments. Pigments other than
chlorophyll that are present in photosyn-
thetic membranes. The accessory pigments
include carotenoids and phycobilins.
acid. A substance that can donate protons. An
acid is converted to its conjugate base by loss
of a proton. (The Lewis theory defines an acid
as an electron-pair acceptor [Lewis acid].)
acid anhydride. The product formed by
condensation of two molecules of acid,
acid dissociation constant (K a ). The equi-
librium constant for the dissociation of a
proton from an acid.

acid-base catalysis. Catalysis in which the
transfer of a proton accelerates a reaction.
ACP. See acyl carrier protein,
activation energy. The free energy required
to promote reactants from the ground state
to the transition state in a chemical reaction,
activator. See transcriptional activator,
active site. The portion of an enzyme that
contains the substrate-binding site and the
amino -acid residues involved in catalyzing
the conversion of substrate(s) to product(s).
Active sites are usually located in clefts be-
tween domains or subunits of proteins or in
indentations on the protein surface,
active transport. The process by which a
solute specifically binds to a transport pro-
tein and is transported across a membrane
against the solute concentration gradient.
Energy is required to drive active transport.
In primary active transport, the energy
source may be light, ATP, or electron trans-
port. Secondary active transport is driven by
ion concentration gradients,
acyl carrier protein (ACP). A protein (in
prokaryotes) or a domain of a protein (in eu-
karyotes) that binds activated intermediates
of fatty acid synthesis via a thioester linkage,
adipocyte. A triacylglycerol-storage cell found
in animals. An adipocyte consists of a fat
droplet surrounded by a thin shell of cytosol in
which the nucleus and other organelles are
suspended.

adipose tissue. Animal tissue composed of
specialized triacylglycerol-storage cells known
as adipocytes.

A-DNA. The conformation of DNA com-
monly observed when purified DNA is dehy-
drated. A-DNA is a right-handed double
helix containing approximately 1 1 base pairs
per turn.

aerobic. Occurring in the presence of oxygen,
affinity chromatography. A chromatographic
technique used to separate a mixture of pro-
teins or other macromolecules in solution
based on specific binding to a ligand that is
covalently attached to the chromatographic
matrix.

affinity labeling. A process by which an en-
zyme (or other macromolecule) is covalently
inhibited by a reaction with a molecule that
specifically interacts with the active site (or
other binding site) .

aldoses. A class of monosaccharides in
which the most oxidized carbon atom, desig-
nated C-l, is aldehydic.
allosteric effector. See allosteric modulator,
allosteric interaction. The modulation of
activity of a protein that occurs when a mole-
cule binds to the regulatory site of the protein,
allosteric modulator. A biomolecule that
binds to the regulatory site of an allosteric
protein and thereby modulates its activity. An
allosteric modulator may be an activator or an
inhibitor. Also known as an allosteric effector,
allosteric protein. A protein whose activity is
modulated by the binding of another molecule,
allosteric site. See regulatory site,
allosteric transitions. The changes in con-
formation of a protein between the active (R)
state and the inactive (T) state.
a helix. A common secondary structure of
proteins, in which the carbonyl oxygen of
each amino acid residue (residue n) forms a
hydrogen bond with the amide hydrogen of
the fourth residue further toward the C-ter-
minus of the polypeptide chain (residue
n + 4). In an ideal right-handed a helix,
equivalent positions recur every 0.54 nm,
each amino acid residue advances the helix by
0.15 nm along the long axis of the helix, and
there are 3.6 amino acid residues per turn,
ammo acid. An organic acid consisting of an
a - carbon atom to which an amino group, a
carboxylate group, a hydrogen atom, and a spe-
cific side chain (R group) are attached. Amino
acids are the building blocks of proteins,
amino acid analysis. A chromatographic
procedure used for the separation and quan-
titation of amino acids in solutions such as
protein hydrolysates,
amino terminus. See N-terminus.
aminoacyl site. See A site

aminoacyl-tRNA synthetase. An enzyme
that catalyzes the activation and attachment
of a specific amino acid to the 3' end of a cor-
responding tRNA molecule,
amphibolic reaction. A metabolic reaction
that can be both catabolic and anabolic,
amphipathic. Describes a molecule that has
both hydrophobic and hydrophilic regions,
amyloplast. Modified chloroplasts that spe-
cialize in starch synthesis,
anabolic reaction. A metabolic reaction that
synthesizes a molecule needed for cell main-
tenance and growth.

anaplerotic reaction. A reaction that re-
plenishes metabolites removed from a central
metabolic pathway (cf. cataplerotic).
angstrom (A). A unit of length equal to
1 X 10 -10 m, or 0.1 nm.
anion. An ion with an overall negative charge,
anode. A positively charged electrode. In elec-
trophoresis, anions move toward the anode,
anomeric carbon. The most oxidized car-
bon atom of a cyclized monosaccharide. The
anomeric carbon has the chemical reactivity
of a carbonyl group.

anomers. Isomers of a sugar molecule that
have different configurations only at the
anomeric carbon atom,
antenna pigments. Light- absorbing pig-
ments associated with the reaction center of a
photosystem. These pigments may form a
separate antenna complex or may be bound
directly to the reaction- center proteins,
antibiotic. A compound, produced by one
organism, that is toxic to other organisms.
Clinically useful antibiotics must be specific
for pathogens and not affect the human host,
antibody. A glycoprotein synthesized by cer-
tain white blood cells as part of the immuno-
logical defense system. Antibodies specifically
bind to foreign compounds, called antigens,
forming antibody- antigen complexes that
mark the antigen for destruction. Also
known as an immunoglobulin,
anticodon. A sequence of three nucleotides
in the anticodon loop of a tRNA molecule.
The anticodon binds to the complementary
codon in mRNA during translation,
anticodon arm. The stem-and-loop struc-
ture in a tRNA molecule that contains the
anticodon.

antigen. A molecule or part of a molecule
that is specifically bound by an antibody,
antiport. The cotransport of two different
species of ions or molecules in opposite direc-
tions across a membrane by a transport protein.

751

752 GLOSSARY OF BIOCHEMICAL TERMS

antisense strand. In double- stranded DNA
the antisense strand is the strand that does
not contain codons. Also called the template
strand. The opposite strand is called the sense
strand or the coding strand,
antisense RNA. An RNA molecule that
binds to a complementary mRNA molecule,
forming a double- stranded region that in-
hibits translation of the mRNA.
apoprotein. A protein whose cofactor(s) is
absent. Without the cofactor(s), the apopro-
tein lacks the biological activity characteristic
of the corresponding holoprotein.
apoptosis. The programed death of a cell,
atomic mass unit. The unit of atomic
weight equal to l/12th the mass of the 12 C
isotope of carbon. The mass of the 12 C nu-
clide is exactly 12 by definition,
attenuation. A mechanism of regulation of
gene expression that couples translation and
transcription. Generally, the translation of a
short reading frame at the beginning of a
prokaryotic operon will determine whether
transcription terminates before the rest of the
operon is transcribed.

autophosphorylation. Phosphorylation of a
protein kinase catalyzed by another molecule
of the same kinase.

autosome. A chromosome other than a sex
chromosome.

autotroph. An organism that can grow and
reproduce using only inorganic substances
(such as C0 2 ) as its only source of essential
elements.

backbone. 1 . The repeating N — C a — C
units connected by peptide bonds in a
polypeptide chain. 2. The repeating sugar-
phosphate units connected by phosphodi-
ester linkages in a nucleic acid,
bacteriophage. A virus that infects a bacter-
ial cell.

base. 1 . A substance that can accept protons.
A base is converted to its conjugate acid by ad-
dition of a proton. (The Lewis theory defines
a base as an electron-pair donor [Lewis
base].) 2. The substituted pyrimidine or
purine of a nucleoside or nucleotide. The het-
erocyclic bases of nucleosides and nucleotides
can participate in hydrogen bonding,
base pairing. The interaction between the
bases of nucleotides in single-stranded nucleic
acids to form double-stranded molecules,
such as DNA, or regions of double-stranded
secondary structure. The most common base
pairs are formed by hydrogen bonding of
adenine (A) with thymine (T) or uracil (U)
and of guanine (G) with cytosine (C).
B-DNA. The most common conformation
of DNA and the one proposed by Watson and
Crick. B-DNA is a right-handed double helix
with a diameter of 2.37 nm and approximately
10.4 base pairs per turn.

P -oxidation pathway. The metabolic path-
way that degrades fatty acids to acetyl CoA,
producing NADH and QH 2 and thereby gen-
erating large amounts of ATR Each round of

P -oxidation of fatty acids consists of four
steps: oxidation, hydration, further oxida-
tion, and thiolysis.
p pleated sheet. See p sheet.

P sheet. A common secondary structure of
proteins that consists of extended polypep-
tide chains stabilized by hydrogen bonds be-
tween the carbonyl oxygen of one peptide
bond and the amide hydrogen of another on
the same or an adjacent polypeptide chain.
The hydrogen bonds are nearly perpendicu-
lar to the extended polypeptide chains, which
may be either parallel (running in the same
N- to C-terminal direction) or antiparallel
(running in opposite directions).

P strand. An extended polypeptide chain
within a P sheet secondary structure or hav-
ing the same conformation as a strand within
a P sheet.

P turn. See turn.

bile. A suspension of bile salts, bile pigments,
and cholesterol that originates in the liver and
is stored in the gall bladder. Bile is secreted
into the small intestine during digestion,
binding-change mechanism. A proposed
mechanism for the phosphorylation of ADP
and release of ATP from FqF! ATP synthase.
The mechanism proposes three different
binding- site conformations for ATP syn-
thase: an open site from which ATP has been
released, an ATP-bearing tight-binding site
that is catalytically active, and an ADP and Pj
loose-binding site that is catalytically inac-
tive. Inward passage of protons through the
ATP synthase complex into the mitochondri-
al matrix causes the open site to become a
loose site; the loose site, already filled with
ADP and Pj, to become a tight site; and the
ATP-bearing site to become an open site,
bioenergetics. The study of energy changes
in biological systems,
biological membrane. See membrane,
biopolymer. A biological macromolecule in
which many identical or similar small mole-
cules are covalently linked to one another to
form a long chain. Proteins, polysaccharides,
and nucleic acids are biopolymers.

Bohr effect. The phenomenon observed when
exposure to carbon dioxide, which lowers the
pH inside the cells, causes the oxygen affinity of
hemoglobin in red blood cells to decrease,
branch migration. The movement of a
crossover, or branch point, resulting in fur-
ther exchange of DNA strands during recom-
bination.

branch site. The point within an intron that
becomes attached to the 5' end of the intron
during splicing of mRNA precursors,
buffer. A solution of an acid and its conju-
gate base that resists changes in pH.
buffer capacity. The ability of a solution to
resist changes in pH. For a given buffer, max-
imum buffer capacity is achieved at the pH at
which the concentrations of the weak acid
and its conjugate base are equal (i.e., when
pH = pKa).

C 4 pathway. A pathway for carbon fixation
in several plant species that minimizes pho-
torespiration by concentrating C0 2 . In this
pathway, C0 2 is incorporated into C 4 acids in
the mesophyll cells, and the C 4 acids are de-
carboxylated in the bundle sheath cells, re-
leasing C0 2 for use by the reductive pentose
phosphate cycle.

calorie (cal). The amount of energy re-
quired to raise the temperature of 1 gram of
water by 1°C (from 14.5°C to 15.5°C). One
calorie is equal to 4.184 J.

Calvin cycle. A cycle of reactions that in-
volve the fixation of carbon dioxide and the
net production of glyceraldehyde-3-phos-
phate. Usually associated with photosynthe-
sis. Also known as the Calvin-Benson cycle,
the C3 pathway, and the reductive pentose
phosphate (RPP) cycle.

Calvin-Benson cycle. See Calvin cycle.

CAM. See Crassulacean acid metabolism,
cap. A 7-methylguanosine residue attached
by a pyrophosphate linkage to the 5' end of a
eukaryotic mRNA molecule. The cap is added
posttranscriptionally and is required for effi-
cient translation. Further covalent modifica-
tions yield alternative cap structures,
carbanion. A carbon anion that results from
the cleavage of a covalent bond between car-
bon and another atom in which both electrons
from the bond remain with the carbon atom,
carbocation. A carbon cation that results
from the cleavage of a covalent bond between
carbon and another atom in which the car-
bon atom loses both electrons from the bond,
carbohydrate. Loosely defined as a com-
pound that is a hydrate of carbon in which the
ratio of C:H:0 is 1:2:1. Carbohydrates include
monomeric sugars (i.e., monosaccharides) and
their polymers. Also known as a saccharide,
carboxyl terminus. See C-terminus.
carnitine shuttle system. A cyclic pathway
that shuttles acetyl CoA from the cytosol to
the mitochondria by formation and trans-
port of acyl carnitine.

cascade. Sequential activation of several
components, resulting in signal amplification,
catabolic reaction. A metabolic reaction
that degrades a molecule to provide smaller
molecular building blocks and energy to an
organism.

catabolite repression. A regulatory mecha-
nism that results in increased rates of tran-
scription of many bacterial genes and
operons when glucose is present. A complex
between cAMP and cAMP regulatory protein
(CRP) activates transcription,
catalytic antibodies. Antibody molecules
that have been genetically manipulated so that
they catalyze reactions involving the antigen,
catalytic center. The polar amino acids in
the active site of an enzyme that participate
in chemical changes during catalysis,
catalytic constant (k cat ). A kinetic constant
that is a measure of how rapidly an enzyme
can catalyze a reaction when saturated with its

GLOSSARY OF BIOCHEMICAL TERMS 753

substrate(s). The catalytic constant is equal to
the maximum velocity ( V max ) divided by the
total concentration of enzyme ([E] tota i), or
the number of moles of substrate converted
to product per mole of enzyme active sites per
second, under saturating conditions. Also
known as the turnover number,
catalytic proficiency. The ratio of the rate
constants for a reaction in the presence of en-
zyme (k cat /K m ) to the rate constant for the
chemical reaction in the absence of enzyme,
cataplerotic reaction. A reaction that re-
moves intermediates in a pathway, especially
the citric acid cycle (cf., anaplerotic) .
cathode. A negatively charged electrode. In
electrophoresis, cations move toward the
cathode.

cation. An ion with an overall positive charge.
cDNA. See complementary DNA.

Central Dogma. The concept that the flow
of information from nucleic acid to protein is
irreversible. The term is often applied incor-
rectly to the actual pathway of information
flow from DNA to RNA to protein,
ceramide. A molecule that consists of a fatty
acid linked to the C-2 amino group of sphin-
gosine by an amide bond. Ceramides are the
metabolic precursors of all sphingolipids.
cerebroside. A glycosphingolipid that con-
tains one monosaccharide residue attached via
a /3-glycosidic linkage to C-l of a ceramide.
Cerebrosides are abundant in nerve tissue and
are found in myelin sheaths,
channel. An integral membrane protein
with a central aqueous passage, which allows
appropriately sized molecules and ions to
traverse the membrane in either direction.
Also known as a pore,
channeling. See metabolite channeling,
chaotropic agent. A substance that enhances
the solubility of nonpolar compounds in
water by disrupting regularities in hydrogen
bonding among water molecules. Concen-
trated solutions of chaotropic agents, such as
urea and guanidinium salts, decrease the hy-
drophobic effect and are thus effective pro-
tein denaturants.

chaperone. A protein that forms complexes
with newly synthesized polypeptide chains
and assists in their correct folding into bio-
logically functional conformations. Chaper-
ones may also prevent the formation of
incorrectly folded intermediates, prevent in-
correct aggregation of unassembled protein
subunits, assist in translocation of polypep-
tide chains across membranes, and assist in
the assembly and disassembly of large multi-
protein structures.

charge-charge interaction. A noncovalent
electrostatic interaction between two charged
particles.

chelate effect. The phenomenon by which the
constant for binding of a ligand having two or
more binding sites to a molecule or atom is
greater than the constant for binding of sepa-
rate ligands to the same molecule or atom.

chemiosmotic theory. A theory proposing
that a proton concentration gradient estab-
lished during oxidation of substrates pro-
vides the energy to drive processes such as the
formation of ATP from ADP and Pj.
chemoautotroph. An autotroph that derives
chemical energy by oxidizing inorganic com-
pounds (cf., photoautotroph),
chemoheterotroph. Non-photosynthetic
organism that requires organic molecules as a
carbon source and derives energy from oxi-
dizing organic molecules,
chemotaxis. A mechanism that couples sig-
nal transduction to flagella movement in
bacteria causing them to move toward a
chemical (positive chemotaxis) or away from
a chemical (negative chemotaxis).
chiral atom. An atom with asymmetric sub-
stitution that can exist in two different con-
figurations.

chloroplast. A chlorophyll- containing or-
ganelle in algae and plant cells that is the site
of photosynthesis.

chromatin. A DNA-protein complex in the
nuclei of eukaryotic cells,
chromatography. A technique used to sepa-
rate components of a mixture based on their
partitioning between a mobile phase, which
can be gas or liquid, and a stationary phase,
which is a liquid or solid,
chromosome. A single DNA molecule con-
taining many genes. An organism may have a
genome consisting of a single chromosome
or many.

chylomicron. A type of plasma lipoprotein
that transports triacylglycerols, cholesterol,
and cholesteryl esters from the small intestine
to the tissues.

citric acid cycle. A metabolic cycle consist-
ing of eight enzyme- catalyzed reactions that
completely oxidizes acetyl units to C0 2 . The
energy released in the oxidation reactions is
conserved as reducing power when the coen-
zymes NAD 1 and ubiquinone (Q) are re-
duced. Oxidation of one molecule of acetyl
CoA by the citric acid cycle generates three
molecules of NADH, one molecule of QH 2 ,
and one molecule of GTP or ATP. Also
known as the Krebs cycle and the tricar-
boxylic acid cycle.

clone. One of the identical copies derived
from the replication or reproduction of a sin-
gle molecule, cell, or organism,
cloning. The generation of many identical
copies of a molecule, cell, or organism.
Cloning sometimes refers to the entire
process of constructing and propagating a re-
combinant DNA molecule,
cloning vector. A DNA molecule that carries
a segment of foreign DNA. A cloning vector
introduces the foreign DNA into a cell where
it can be replicated and sometimes expressed,
coding strand. The strand of DNA within a
gene whose nucleotide sequence is identical
to that of the RNA produced by transcription
(with the replacement of T by U in RNA).

codon. A sequence of three nucleotide
residues in mRNA (or DNA) that specifies a
particular amino acid according to the ge-
netic code.

coenzyme. An organic molecule required by
an enzyme for full activity. Coenzymes can be
further classified as cosubstrates or prosthetic
groups.

coenzyme A. A large coenyme used in trans-
ferring acyl groups.

cofactor. An inorganic ion or organic mole-
cule required by an apoenzyme to convert it
to a holoenzyme. There are two types of co-
factors: essential ions and coenzymes,
column chromatography. A technique for
purifying proteins. See affinity chromatog-
raphy, gel-filtration chromatography, ion-
exchange chromatography, HPLC, and affinity
chromatography.

competitive inhibition. Reversible inhibi-
tion of an enzyme- catalyzed reaction by an
inhibitor that prevents substrate binding,
complementary DNA (cDNA). DNA syn-
thesized from an mRNA template by the ac-
tion of reverse transcriptase,
concerted theory of cooperativity and al-
losteric regulation. A model of the coopera-
tive binding of ligands to oligomeric
proteins. According to the concerted theory,
the change in conformation of a protein due
to the binding of a substrate or an allosteric
modulator shifts the equilibrium of the con-
formation of the protein between T (a low
substrate- affinity conformation) and R (a
high substrate-affinity conformation). This
theory suggests that all subunits of the pro-
tein have the same conformation, either all T
or all R. Also known as the symmetry- driven
theory.

condensation. A reaction involving the join-
ing of two or more molecules accompanied
by the elimination of water, alcohol, or other
simple substance.

configuration. A spatial arrangement of
atoms that cannot be altered without break-
ing and re-forming covalent bonds,
conformation. Any three-dimensional struc-
ture, or spatial arrangement, of a molecule
that results from rotation of functional groups
around single bonds. Because there is free ro-
tation around single bonds, a molecule can
potentially assume many conformations,
conjugate acid. The product resulting from
the gain of a proton by a base,
conjugate base. The product resulting from
the loss of a proton by an acid,
consensus sequence. The sequence of nu-
cleotides most commonly found at each posi-
tion within a region of DNA or RNA.
cooperativity. 1 . The phenomenon whereby
the binding of one ligand or substrate mole-
cule to a protein influences the affinity of the
protein for additional molecules of the same
substance. Cooperativity may be positive or
negative. 2. The phenomenon whereby formation

754 GLOSSARY OF BIOCHEMICAL TERMS

of structure in one part of a macromolecule
promotes the formation of structure in the
rest of the molecule.

core particle. See nucleosome core particle,
corepressor. A ligand that binds to a repres-
sor of a gene causing it to bind DNA and pre-
vent transcription.

Cori cycle. An interorgan metabolic loop that
recycles carbon and transports energy from
the liver to the peripheral tissues. Glucose is
released from the liver and metabolized to
produce ATP in other tissues. The resulting
lactate is then returned to the liver for conver-
sion back to glucose by gluconeogenesis.
cosubstrate. A coenzyme that is a substrate
in an enzyme-catalyzed reaction. A cosub-
strate is altered during the course of the reac-
tion and dissociates from the active site of the
enzyme. The original form of the cosubstrate
can be regenerated in a subsequent enzyme-
catalyzed reaction.

cotransport. The coupled transport of two
different species of solutes across a mem-
brane, in the same direction (symport) or the
opposite direction (antiport), carried out by
a transport protein.

coupled reactions. Two metabolic reactions
that share a common intermediate,
covalent catalysis. Catalysis in which one
substrate, or part of it, forms a covalent bond
with the catalyst and then is transferred to a
second substrate. Many enzymatic group-
transfer reactions proceed by covalent catalysis.
Crassulacean acid metabolism (CAM). A
modified sequence of carbon-assimilation
reactions used primarily by plants in arid en-
vironments to reduce water loss during pho-
tosynthesis. In these reactions, C0 2 is taken
up at night, resulting in the formation of
malate. During the day, malate is decarboxy-
lated, releasing C0 2 for use by the reductive
pentose phosphate cycle.

C-terminus. The amino acid residue bearing
a free carboxyl group at one end of a peptide
chain. Also known as the carboxyl terminus,
cyclic electron transport. A modified se-
quence of electron transport steps in chloro-
plasts that operates to provide ATP without
the simultaneous formation of NADPH.
cytoplasm. The part of a cell enclosed by the
plasma membrane, excluding the nucleus,
cytoskeleton. A network of proteins that
contributes to the structure and organization
of a eukaryotic cell.

cytosol. The aqueous portion of the cyto-
plasm minus the subcellular structures.

D arm. The stem-and-loop structure in a
tRNA molecule that contains dihydrouridy-
late (d) residues.

dalton. A unit of mass equal to one atomic
mass unit.

dark reactions. The photosynthetic reac-
tions in which NADPH and ATP are used to
fix C0 2 to carbohydrate. Also known as the
light-independent reactions.

degeneracy. When referring to the genetic
code, degeneracy refers to the fact that several
different codons specify the same amino acid,
dehydrogenase. An enzyme that catalyzes
the removal of hydrogen from a substrate or
the oxidation of a substrate. Dehydrogenases
are members of the IUBMB class of enzymes
known as oxidoreductases.
denaturation. 1 . A disruption in the native
conformation of a biological macromolecule
that results in loss of the biological activity of
the macromolecule. 2. The complete un-
winding and separation of complementary
strands of DNA.

detergent. An amphipathic molecule consist-
ing of a hydrophobic portion and a hydrophilic
end that may be ionic or polar. Detergent mol-
ecules can aggregate in aqueous media to form
micelles. Also known as a surfactant,
dialysis. A procedure in which low-molecu-
lar- weight solutes in a sample are removed by
diffusion through a semipermeable barrier
and replaced by solutes from the surrounding
medium.

diffusion controlled reaction. A reaction that
occurs with every collision between reactant
molecules. In enzyme-catalyzed reactions,
the k CSLt /K m ratio approaches a value of
10 8 - 10 9 M _1 s _1 .

diploid. Having two sets of chromosomes or
two copies of the genome,
dipole. Two equal but opposite charges, sep-
arated in space, resulting from the uneven
distribution of charge within a molecule or a
chemical bond.

direct repair. The removal of DNA damage by
proteins that recognize damaged nucleotides
and mismatched bases and repair them with-
out cleaving the DNA or excising the base,
distributive enzyme. An enzyme that dissoci-
ates from its growing polymeric product after
addition of each monomeric unit and must re-
associate with the polymer for polymerization
to proceed (c£, progressive enzyme),
disulfide bond. A covalent linkage formed
by oxidation of the sulfhydryl groups of two
cysteine residues. Disulfide bonds are impor-
tant in stabilizing the three-dimensional
structures of some proteins,
domain. A discrete, independent folding
unit within the tertiary structure of a protein.
Domains are usually combinations of several
motifs forming a characteristic fold.

double helix. A nucleic acid conformation
in which two antiparallel polynucleotide
strands wrap around each other to form a
two- stranded helical structure stabilized
largely by stacking interactions between adja-
cent hydrogen-bonded base pairs.

double-reciprocal plot. A plot of the recip-
rocal of initial velocity versus the reciprocal
of substrate concentration for an enzyme-
catalyzed reaction. The x and y intercepts
indicate the values of the reciprocals of the
Michaelis constant and the maximum velocity,

respectively. A double-reciprocal plot is a lin-
ear transformation of the Michaelis-Menten
equation. Also known as a Lineweaver-
Burk plot.

E. See reduction potential.

E or . See standard reduction potential.

E site. Exit site. The site on a ribosome from
which a deaminoacylated tRNA is released
during protein synthesis.

Edman degradation. A procedure used to
determine the sequence of amino acid
residues from a free N-terminus of a
polypeptide chain. The N - terminal residue is
chemically modified, cleaved from the chain,
and identified by chromatographic proce-
dures, and the rest of the polypeptide is
recovered. Multiple reaction cycles allow
identification of the new N-terminal residue
generated by each cleavage step,
effector enzyme. A membrane-associated
protein that produces an intracellular second
messenger in response to a signal from a
transducer.

eicosanoid. An oxygenated derivative of a
20-carbon polyunsaturated fatty acid.
Eicosanoids function as short-range messen-
gers in the regulation of various physiological
processes.

electromotive force (emf). A measure of the
difference between the reduction potentials
of the reactions on the two sides of an elec-
trochemical cell (i.e., the voltage difference
produced by the reactions) .
electrolyte. A molecule such as NaCl that
can dissociated to form ions.

electron transport. A set of reactions in
which compounds such as NADH and re-
duced ubiquinone (QH 2 ) are aerobically oxi-
dized and ATP is generated from ADP and Pj.
Membrane- associated electron transport
consists of two tightly coupled phenomena:
oxidation of substrates by the respiratory
electron transport chain, accompanied by the
translocation of protons across the inner mi-
tochondrial membrane to generate a proton
concentration gradient; and formation of
ATP, driven by the flux of protons into the
matrix through a channel in ATP synthase.

electrophile. A positively charged or electron-
deficient species that is attracted to chemical
species that are negatively charged or contain
unshared electron pairs (nucleophiles).

electrophoresis. A technique used to sepa-
rate molecules by their migration in an elec-
tric field, primarily on the basis of their net
charge.

electrospray mass spectrometry. A tech-
nique in mass spectrometry where the target
molecule is sprayed into the detector in tiny
droplets.

electrostatic interaction. A general term for
the electronic interaction between particles.
Electrostatic interactions include charge-
charge interactions, hydrogen bonds, and van
der Waals forces.

GLOSSARY OF BIOCHEMICAL TERMS 755

elongation factor. A protein that is involved
in extending the peptide chain during pro-
tein synthesis.

enantiomers. Stereoisomers that are non-
superimposable mirror images,
endocytosis. The process by which matter is
engulfed by a plasma membrane and brought
into the cell within a lipid vesicle derived
from the membrane.

endonuclease. An enzyme that catalyzes the
hydrolysis of phosphodiester linkages at vari-
ous sites within polynucleotide chains,
endoplasmic reticulum. A membranous
network of tubules and sheets continuous
with the outer nuclear membrane of eukary-
otic cells. Regions of the endoplasmic reticu-
lum coated with ribosomes are called the
rough endoplasmic reticulum; regions hav-
ing no attached ribosomes are known as the
smooth endoplasmic reticulum. The endo-
plasmic reticulum is involved in the sorting
and transport of certain proteins and in the
synthesis of lipids.

endosomes. Smooth vesicles inside the cell
that are receptacles for endocytosed material,
energy-rich compound. A compound whose
hydrolysis occurs with a large negative free-
energy change (equal to or greater than that
for ATP — » ADP + PJ.

enthalpy (H). A thermodynamic state func-
tion that describes the heat content of a system,
entropy (S). A thermodynamic state func-
tion that describes the randomness or disor-
der of a system.

enzymatic reaction. A reaction catalyzed by
a biological catalyst, an enzyme. Enzymatic
reactions are 10 3 to 10 17 times faster than the
corresponding uncatalyzed reactions,
enzyme. A biological catalyst, almost always
a protein. Some enzymes may require addi-
tional cofactors for activity. Virtually all bio-
chemical reactions are catalyzed by specific
enzymes.

enzyme assay. A method used to analyze the
activity of a sample of an enzyme. Typically,
enzymatic activity is measured under selected
conditions such that the rate of conversion of
substrate to product is proportional to en-
zyme concentration.

enzyme inhibitor. A compound that binds
to an enzyme and interferes with its activity
by preventing either the formation of the ES
complex or its conversion to E + P.
enzyme-substrate complex (ES). A complex
formed when substrate molecules bind non-
covalently within the active site of an
enzyme.

epimers. Isomers that differ in configuration
at only one of several chiral centers,
equilibrium. The state of a system in which
the rate of conversion of substrate to product
is equal to the rate of conversion of product
to substrate. The free-energy change for a re-
action or system at equilibrium is zero.

equilibrium constant (K e q ). The ratio of the
concentrations of products to the concentra-
tions of reactants at equilibrium. The equi-
librium constant is related to the standard
Gibbs free energy change of reaction,
essential amino acid. An amino acid that
cannot be synthesized by an animal and must
be obtained in the diet,
essential fatty acid. A fatty acid that cannot
be synthesized by an animal and must be ob-
tained in the diet.

essential ion. An ion required as a cofactor
for the catalytic activity of certain enzymes.
Some essential ions, called activator ions, are
reversibly bound to enzymes and often par-
ticipate in the binding of substrates, whereas
tightly bound metal ions frequently partici-
pate directly in catalytic reactions,
eukaryote. An organism whose cells gener-
ally possess a nucleus and internal mem-
branes (cf., prokaryote),
excision repair. The reversal of DNA dam-
age by excision- repair endonucleases. Gross
lesions that alter the structure of the DNA
helix are repaired by cleavage on each side of
the lesion and removal of the damaged DNA.
The resulting single- stranded gap is filled by
DNA polymerase and sealed by DNA ligase.
exocytosis. The process by which material
destined for secretion from a cell is enclosed
in lipid vesicles that are transported to and
fuse with the plasma membrane, releasing
the material into the extracellular space,
exon. A nucleotide sequence that is present
in the primary RNA transcript and in the
mature RNA molecule. The term exon also
refers to the region of the gene that corre-
sponds to a sequence present in the mature
RNA (cf., intron).

exonuclease. An enzyme that catalyzes the
sequential hydrolysis of phosphodiester link-
ages from one end of a polynucleotide chain,
extrinsic membrane protein. See peripheral
membrane protein.

facilitated diffusion. See passive transport,
facultative anaerobe. An organism that can
survive in the presence or absence of oxygen,
fatty acid. A long chain aliphatic hydrocar-
bon with a single carboxyl group at one end.
Fatty acids are the simplest type of lipid and
are components of many more complex
lipids, including triacylglycerols, glycerophos-
pholipids, sphingolipids, and waxes,
feedback inhibition. Inhibition of an en-
zyme that catalyzes an early step in a meta-
bolic pathway by an end product of the same
pathway.

feed-forward activation. Activation of an
enzyme in a metabolic pathway by a metabo-
lite produced earlier in the pathway,
fermentation. The anaerobic catabolism of
metabolites for energy production. In alco-
holic fermentation, pyruvate is converted to
ethanol and carbon dioxide.

fibrous proteins. A major class of water-in-
soluble proteins that associate to form long
fibers. Many fibrous proteins are physically
tough and provide mechanical support to in-
dividual cells or entire organisms,
first-order reaction. A reaction whose rate
is directly proportional to the concentration
of only one reactant.

Fischer projection. A two-dimensional rep-
resentation of the three-dimensional struc-
tures of sugars and related compounds. In a
Fischer projection, the carbon skeleton is
drawn vertically, with C- 1 at the top. At a chi-
ral center, horizontal bonds extend toward
the viewer and vertical bonds extend away
from the viewer.

fluid mosaic model. A model proposed for
the structure of biological membranes. In
this model, the membrane is depicted as a dy-
namic structure in which lipids and mem-
brane proteins (both integral and peripheral)
rotate and undergo lateral diffusion,
fluorescence. A form of luminescence in
which visible radiation is emitted from a
molecule as it passes from a higher to a lower
electronic state.

flux. The flow of material through a meta-
bolic pathway. Flux depends on the supply of
substrates, the removal of products, and the
catalytic capabilities of the enzymes involved
in the pathway.

fold. A combination of secondary structures
that form the core of a protein domain. Many
different folds have been characterized,
frameshift mutation. An alteration in DNA
caused by the insertion or deletion of a num-
ber of nucleotides not divisible by three. A
frameshift mutation changes the reading
frame of the corresponding mRNA molecule
and affects translation of all codons down-
stream of the mutation,
free energy change. See Gibbs free energy
change.

free radical. A molecule or atom with an
unpaired electron.

furanose. A monosaccharide structure that
forms a five-membered ring as a result of in-
tramolecular hemiacetal formation.

G protein. A protein that binds guanine nu-
cleotides.

AG. Sec Gibbs free energy change.

A G ° ' . See standard Gibbs free energy change,
ganglioside. A glycosphingolipid in which
oligosaccharide chains containing AT-acetyl-
neuraminic acid are attached to a ceramide.
Gangliosides are present on cell surfaces and
provide cells with distinguishing surface
markers that may serve in cellular recogni-
tion and cell-to-cell communication,
gas chromatography. A chromatographic
technique used to separate components of a
mixture based on their partitioning between
the gas phase and a stationary phase, which
can be a liquid or solid.

756 GLOSSARY OF BIOCHEMICAL TERMS

gel-filtration chromatography. A chromato-
graphic technique used to separate a mixture
of proteins or other macromolecules in solu-
tion based on molecular size, using a matrix of
porous beads. Also known as molecular- exclu-
sion chromatography.

gene. Loosely defined as a segment of DNA
that is transcribed. In some cases, the term
gene may also be used to refer to a segment of
DNA that encodes a functional protein or
corresponds to a mature RNA molecule,
genetic code. The correspondence between
a particular three nucleotide codon and the
amino acid it specifies. The standard genetic
code of 64 codons is used by almost all or-
ganisms. The genetic code is used to translate
the sequence of nucleotides in mRNA into
protein.

genetic recombination. The exchange or
transfer of DNA from one molecule of DNA
to another (cf., homologous recombination),
genome. One complete set of the genetic in-
formation in an organism. It may be a single
chromosome or a set of chromosomes (hap-
loid). Mitochondria and chloroplasts have
genomes separate from that in the nucleus of
eukaryotic cells.

Gibbs free energy change (AG). A thermo-
dynamic quantity that defines the equilib-
rium condition in terms of the changes in
enthalpy ( H ) and entropy (S) of a system at
constant pressure. AG = A H — TAS, where
T is absolute temperature. Free energy is a
measure of the energy available within a sys-
tem to do work.

globular proteins. A major class of proteins,
many of which are water soluble. Globular
proteins are compact and roughly spherical,
containing tightly folded polypeptide chains.
Typically, globular proteins include indenta-
tions, or clefts that specifically recognize and
transiently bind other compounds,
glucogenic compound. A compound, such
as an amino acid, that can be used for gluco-
neogenesis in animals.

gluconeogenesis. A pathway for synthesis of
glucose from a noncarbohydrate precursor.
Gluconeogenesis from pyruvate involves the
seven near- equilibrium reactions of glycoly-
sis traversed in the reverse direction. The
three metabolically irreversible reactions of
glycolysis are bypassed by four enzymatic re-
actions that do not occur in glycolysis,
glucoside. A glycoside where the anomeric
carbon atom is from glucose.

glycan. A general term for an oligosaccharide
or a polysaccharide. A homoglycan is a poly-
mer of identical monosaccharide residues; a
heteroglycan is a polymer of different mono-
saccharide residues.

glycerophospholipid. A lipid consisting of
two fatty acyl groups bound to C-l and C-2 of
glycerol 3 -phosphate and, in most cases, a
polar substituent attached to the phosphate
moiety. Glycerophospholipids are major com-
ponents of biological membranes.

glycoconjugate. A carbohydrate derivative
in which one or more carbohydrate chains
are covalently linked to a peptide chain, pro-
tein, or lipid.

glycoforms. Glycoproteins containing iden-
tical amino acid sequences but different
oligosaccharide-chain compositions.

glycogen. A branched homopolymer of glu-
cose residues joined by a-(l — >4) linkages
with a-( 1 — > 6) linkages at branch points.
Glycogen is a storage polysaccharide in ani-
mals and bacteria.

glycolysis. A catabolic pathway consisting of
10 enzyme-catalyzed reactions by which one
molecule of glucose is converted to two mole-
cules of pyruvate. In the process, two molecules
of ATP are formed from ADP + P*, and two
molecules of NAD 1 are reduced to NADH.

glycoprotein. A protein that contains cova-
lently bound carbohydrate residues.

glycosaminoglycan. An unbranched poly-
saccharide of repeating disaccharide units.
One component of the disaccharide is an
amino sugar; the other component is usually
a uronic acid.

glycoside. A molecule containing a carbohy-
drate in which the hydroxyl group of the
anomeric carbon has been replaced through
condensation with an alcohol, an amine, or a
thiol.

glycosidic bond. Acetal linkage formed by
condensation of the anomeric carbon atom
of a saccharide with a hydroxyl, amino, or
thiol group of another molecule. The most
commonly encountered glycosidic bonds are
formed between the anomeric carbon of one
sugar and a hydroxyl group of another sugar.
Nucleosidic bonds are N-linked glycosidic
bonds.

glycosphingolipid. A lipid containing
sphingosine and carbohydrate moieties.

glycosylation. See protein glycosylation.

glyoxylate cycle. A variation of the citric
acid cycle in certain plants, bacteria, and
yeast that allows net production of glucose
from acetyl CoA via oxaloacetate. The gly-
oxylate cycle bypasses the two C0 2 2 produc-
ing steps of the citric acid cycle.

glyoxysome. An organelle that contains spe-
cialized enzymes for the glyoxylate cycle.

Golgi apparatus. A complex of flattened,
fluid- filled membranous sacs in eukaryotic
cells, often found in proximity to the endo-
plasmic reticulum. The Golgi apparatus is
involved in the modification, sorting, and
targeting of proteins.

granum. A stack of flattened vesicles formed
from the thylakoid membrane in chloroplasts.

group transfer potential. See photsphoryl
group transfer potential.

group transfer reaction. A reaction in
which a substituent or functional group is
transferred from one substrate to another.

H. See enthalpy.

hairpin. 1 . A secondary structure adopted by
single- stranded polynucleotides that arises
when short regions fold back on themselves
and hydrogen bonds form between comple-
mentary bases. Also known as a stem-loop.
2. A tight turn connecting two consecutive
/3 strands of a polypeptide,
haploid. Having one set of chromosomes or
one copy of the genome (cf., diploid),
high energy molecule. See energy-rich
compound.

Haworth projection. A representation in
which a cyclic sugar molecule is depicted as a
flat ring that is projected perpendicular to the
plane of the page. Heavy lines represent the
part of the molecule that extends toward the
viewer.

HDL. See high density lipoprotein,
heat of vaporization. The amount of heat
required to evaporate 1 gram of a liquid,
heat shock protein. A protein whose synthe-
sis is increased in response to stresses such as
high temperature. Many heat shock proteins
are chaperones that are also expressed in the
absence of stress.

helicase. An enzyme that is involved in un-
winding DNA.

hemiacetal. The product formed when an
alcohol reacts with an aldehyde,
hemiketal. The product formed when an al-
cohol reacts with a ketone.
Henderson-Hasselbalch equation. An equa-
tion that describes the pH of a solution of a
weak acid or a weak base in terms of the p K a
and the concentrations of the proton donor
and proton acceptor forms,
heterochromatin. Regions of chromatin
that are highly condensed,
heterocyclic molecule. A molecule that con-
tains a ring structure made up of more than
one type of atom.

heteroglycan (heteropolysaccharide). A car-
bohydrate polymer whose residues consist of
two or more different types of monosaccharide,
heterotroph. An organism that requires at
least one organic nutrient, such as glucose, as
a carbon source.

high density lipoprotein (HDL). A type of
plasma lipoprotein that is enriched in protein
and transports cholesterol and cholesteryl es-
ters from tissues to the liver,
high-performance liquid chromatography
(HPLC). A chromatographic technique used
to separate components of a mixture by
dissolving the mixture in a liquid solvent and
forcing it to flow through a chromatographic
column under high pressure,
histones. A class of proteins that bind to
DNA to form chromatin. The nuclei of eu-
karyotic cells contain five histones, known as
HI, H2A, H2B, H3, and H4.

Holliday junction. The region of strand
crossover resulting from recombination be-
tween two molecules of homologous double-
stranded DNA.

GLOSSARY OF BIOCHEMICAL TERMS 757

homoglycan (homopolysaccharide). A car-
bohydrate polymer whose residues consist of
a single type of monosaccharide,
homologous. Referring to genes or proteins
that descend from a common ancestor,
homologous recombination. Recombination
between molecules of DNA that have closely
related sequences (i.e., they are homologous).
This is the standard form of recombination
that occurs between chromosomes in eukary-
otic cells.

homology. The similarity of genes or pro-
teins as a result of evolution from a common
ancestor.

hormone response element. A DNA se-
quence that binds a transcriptional activator
consisting of a steroid hormone receptor
complex.

housekeeping genes. Genes that encode
proteins or RNA molecules that are essential
for the normal activities of all living cells.
HPLC. See high-performance liquid chro-
matography.

hydration. A state in which a molecule or
ion is surrounded by water,
hydrogen bond. A weak electrostatic interac-
tion the formed when a hydrogen atom bonded
covalently to a strongly electronegative atom
is partially shared by interacting with electron
pair of another electronegative atom,
hydrolase. An enzyme that catalyzes the
hydrolytic cleavage of its substrate(s) (i.e.,
hydrolysis).

hydropathy. A measure of the hydrophobic-
ity of amino acid side chains. The more posi-
tive the hydropathy value, the greater the
hydrophobicity.

hydrophilic. “Water loving” — describing

molecules that interact favorably with water,
hydrophilicity. The degree to which a com-
pound or functional group interacts with
water or is preferentially soluble in water,
hydrophobic. “Water fearing” — describing
molecules that do not interact favorably with
water and are much less soluble than hy-
drophilic molecules.

hydrophobic effect. The exclusion of hy-
drophobic groups or molecules by water. The
hydrophobic effect appears to depend on the
increase in entropy of solvent water mole-
cules that are released from an ordered
arrangement around the hydrophobic group,
hydrophobic interaction. A weak, noncova-
lent interaction between nonpolar molecules
or substituents that results from the strong
association of water molecules with one an-
other. Such association leads to the shielding
or exclusion of nonpolar molecules from an
aqueous environment.

hydrophobicity. The degree to which a
compound or functional group that is solu-
ble in nonpolar solvents is insoluble or only
sparingly soluble in water.

IDL. See intermediate density lipoprotein,
induced fit. Activation of an enzyme by a

substrate- initiated conformational change,
inducer. A ligand that binds to and inactivates
a repressor thereby increasing the transcription
of the gene controlled by the repressor,
inhibition constant (K{). The equilibrium
constant for the dissociation of an inhibitor
from an enzyme-inhibitor complex,
inhibitor. A compound that binds to an en-
zyme and inhibits its activity
initial velocity (v 0 )* The rate of conversion
of substrate to product in the early stages of
an enzymatic reaction, before appreciable
product has been formed,
initiation codon. A codon that specifies the
initiation site for protein synthesis. The me-
thionine codon (AUG) is the most common
initiation codon.

initiation factor. See translation initiation
factor.

initiator tRNA. The tRNA molecule that is
used exclusively at initiation codons. The
initiator tRNA is usually a specific me-
thionyl-tRNA.

integral membrane protein. A membrane
protein that penetrates the hydrophobic core
of the lipid bilayer and usually spans the bi-
layer completely Also known as an intrinsic
membrane protein.

intercalating agent. A compound contain-
ing a planar ring structure that can fit be-
tween the stacked base pairs of DNA.
Intercalating agents distort the DNA struc-
ture, partially unwinding the double helix,
intermediary metabolism. The metabolic
reactions by which the small molecules of
cells are interconverted.
intermediate density lipoprotein (IDL). A
type of plasma lipoprotein that is formed
during the breakdown of VLDLs.
intermediate filament. A structure com-
posed of different protein subunits, found in
the cytoplasm of most eukaryotic cells. Inter-
mediate filaments are components of the cy-
toskeletal network.

intron. An internal nucleotide sequence that
is removed from the primary RNA transcript
during processing. The term intron also
refers to the region of the gene that corre-
sponds to the corresponding RNA intron
(cf., exon).

inverted repeat. A sequence of nucleotides
that is repeated in the opposite orientation
within the same polynucleotide strand. An
inverted repeat in double- stranded DNA can
give rise to a cruciform structure,
ion pair. An electrostatic interaction between
ionic groups of opposite charge within the in-
terior of a macromolecule such as a globular
protein.

ion product for water (1C W ). The product of
the concentrations of hydronium ions and
hydroxide ions in an aqueous solution, equal
to 1.0 X 1(T 14 M 2 .

ion-exchange chromatography. A chromato-
graphic technique used to separate a mixture of

ionic species in solution, using a charged ma-
trix. In anion-exchange chromatography, a
positively charged matrix binds negatively
charged solutes, and in cation-exchange chro-
matography, a negatively charged matrix binds
positively charged solutes. The bound species
can be serially eluted from the matrix by grad-
ually changing the pH or increasing the salt
concentration in the solvent,
ionophore. A compound that facilitates the
diffusion of ions across bilayers and mem-
branes by serving as a mobile ion carrier or
by forming a channel for ion passage,
irreversible enzyme inhibition. A form of
enzyme inhibition where the inhibitor binds
covalently to the enzyme,
isoacceptor tRNA molecules. Different tRNA
molecules that bind the same amino acid,
isoelectric focusing. A modified form of
electrophoresis that uses buffers to create a
pH gradient within a polyacrylamide gel.
Each protein migrates to its isoelectric point
(pi), that is, the pH in the gradient at which it
no longer carries a net positive or negative
charge.

isoelectric point (pi). The pH at which a
zwitterionic molecule does not migrate in an
electric field because its net charge is zero,
isoenzymes. See isozymes,
isomerase. An enzyme that catalyzes an iso-
merization reaction, a change in geometry or
structure within one molecule,
isoprene. A branched, unsaturated five-car-
bon molecule that forms the basic structural
unit of all isoprenoids, including the steroids
and lipid vitamins.

isoprenoid. A lipid that is structurally related
to isoprene.

isozymes. Different proteins from a single
biological species that catalyze the same reac-
tion. Also known as isoenzymes,
junk DNA. Regions of the genome with no
known function.

K a . See acid dissociation constant,
kb. See kilobase pair.
k cat . See catalytic constant.

^cat / K m . The second-order rate constant
for conversion of enzyme and substrate to
enzyme and product at low substrate con-
centrations. The ratio of k CdX to iC m , when
used to compare several substrates, is called
the specificity constant.

K eq . See equilibrium constant,
ketogenesis. The pathway that synthesizes
ketone bodies from acetyl CoA in the mito-
chondrial matrix in mammals,
ketogenic compound. A compound, such as
an amino acid, that can be degraded to form
acetyl CoA and can thereby contribute to the
synthesis of fatty acids or ketone bodies,
ketone bodies. Small molecules that are
synthesized in the liver from acetyl CoA.
During starvation, the ketone bodies
/3 -hydroxybutyrate and acetoacetate become
major metabolic fuels.

758 GLOSSARY OF BIOCHEMICAL TERMS

ketoses. A class of monosaccharides in which
the most oxidized carbon atom, usually C-2,
is ketonic.

Ky See inhibition constant,
kilobase pair (kb). A unit of length of dou-
ble-stranded DNA, equivalent to 1000 base
pairs.

kinase. An enzyme that catalyzes transfer of
a phosphoryl group to an acceptor molecule.
A protein kinase catalyzes the phosphoryla-
tion of protein substrates. Kinases are also
known as phosphotransferases,
kinetic mechanism. A scheme used to de-
scribe the sequence of steps in a multisub-
strate enzyme-catalyzed reaction,
kinetic order. The sum of the exponents in a
rate equation, which reflects how many mol-
ecules are reacting in the slowest step of the
reaction. Also known as reaction order.

K m . See Michaelis constant.

Krebs cycle. See citric acid cycle.

K w . See ion product of water,
lagging strand. The newly synthesized DNA
strand formed by discontinuous 5' — > 3'
polymerization in the direction opposite
replication fork movement,
lateral diffusion. The rapid motion of lipid
or protein molecules within the plane of one
leaflet of a lipid bilayer.

LDL. See low density lipoprotein,
leader peptide. The peptide encoded by a por-
tion of the leader region of certain regulated
operons. Synthesis of a leader peptide is the
basis for regulating transcription of the entire
operon by the mechanism of attenuation,
leader region. The sequence of nucleotides
that lie between the transcription start site
and the first coding region of an operon.
leading strand. The newly synthesized DNA
strand formed by continuous 5' — > 3' poly-
merization in the same direction as replica-
tion fork movement,
leaflet. One layer of a lipid bilayer,
lectin. A plant protein that binds specific
saccharides in glycoproteins,
leucine zipper. A structural motif found in
DNA-binding proteins and other proteins.
The zipper is formed when the hydrophobic
faces (frequently containing leucine residues)
of two amphipathic a -helices from the same
or different polypeptide chains interact to
form a coiled-coil structure.

LHC. See light-harvesting complex,
ligand. A molecule, group, or ion that binds
noncovalently to another molecule or atom,
ligand-gated ion channel. A membrane ion
channel that opens or closes in response to
binding of a specific ligand,
ligase. An enzyme that catalyzes the joining,
or ligation, of two substrates. Ligation reac-
tions require the input of the chemical poten-
tial energy of a nucleoside triphosphate such
as ATP. Ligases are commonly referred to as
synthetases.

light reactions. The photosynthetic reac-
tions in which protons derived from water
are used in the chemiosmotic synthesis of
ATP from ADP + Pj and a hydride ion from
water reduces to NADPH. Also known as the
light-dependent reactions,
light-harvesting complex (LHC). A large
pigment complex in the thylakoid membrane
that aids a photosystem in gathering light,
limit dextrin. A branched oligosaccharide
derived from a glucose polysaccharide by the
hydrolytic action of amylase or the phospho-
rolytic action of glycogen phosphorylase or
starch phosphorylase. Limit dextrins are re-
sistant to further degradation catalyzed by
amylase or phosphorylase. Limit dextrins can
be further degraded only after hydrolysis of
the a-(l — » 6) linkages.

Lineweaver-Burk plot. See double-reciprocal
plot.

linker DNA. The stretch of DNA (approxi-
mately 54 base pairs) between two adjacent
nucleosome core particles,
lipase. An enzyme that catalyzes the hydrol-
ysis of triacylglycerols.

lipid. A water- insoluble (or sparingly solu-
ble) organic compound found in biological
systems, which can be extracted by using rel-
atively nonpolar organic solvents,
lipid bilayer. A double layer of lipids in
which the hydrophobic tails associate with
one another in the interior of the bilayer and
the polar head groups face outward into the
aqueous environment. Lipid bilayers are the
structural basis of biological membranes,
lipid raft. A patch of membrane rich in cho-
lesterol and sphingolipid.
lipid vitamin. A polyprenyl compound com-
posed primarily of a long hydrocarbon chain
or fused ring. Unlike water-soluble vitamins,
lipid vitamins can be stored by animals. Lipid
vitamins include vitamins A, D, E, and K.
lipid anchored membrane protein. A mem-
brane protein that is tethered to a mem-
brane through covalent linkage to a lipid
molecule.

lipopolysaccharide. A macromolecule com-
posed of lipid A (a disaccharide of phosphory-
lated glucosamine residues with attached fatty
acids) and a polysaccharide. Lipopolysaccha-
rides are found in the outer membrane of
gram-negative bacteria. These compounds are
released from bacteria undergoing lysis and
are toxic to humans and other animals. Also
known as an endotoxin,
lipoprotein. A macromolecular assembly of
lipid and protein molecules with a hydropho-
bic core and a hydrophilic surface. Lipids are
transported via lipoproteins,
liposome. A synthetic vesicle composed of a
phospholipid bilayer that encloses an aque-
ous compartment.

loop. A nonrepetitive polypeptide region
that connects secondary structures within a
protein molecule and provides directional

changes necessary for a globular protein to
attain its compact shape. Loops contain from
2 to 16 residues. Short loops of up to 5
residues are often called turns,
low density lipoprotein (LDL). A type of
plasma lipoprotein that is formed during the
breakdown of IDLs and is enriched in choles-
terol and cholesteryl esters,
lumen. The aqueous space enclosed by a bi-
ological membrane, such as the membrane of
the endoplasmic reticulum or the thylakoid
membrane.

lyase. An enzyme that catalyzes a nonhy-
drolytic or nonoxidative elimination reaction,
or lysis, of a substrate, with the generation of a
double bond. In the reverse direction, a lyase
catalyzes addition of one substrate to a double
bond of a second substrate,
lysophosphoglyceride. An amphipathic
lipid that is produced when one of the two
fatty acyl moieties of a glycerophospholipid is
hydrolytically removed. Low concentrations
of lysophosphoglycerides are metabolic in-
termediates, whereas high concentrations
disrupt membranes, causing cells to lyse,
lysosome. A specialized digestive organelle
in eukaryotic cells. Lysosomes contain a vari-
ety of enzymes that catalyze the breakdown
of cellular biopolymers, such as proteins, nu-
cleic acids, and polysaccharides, and the di-
gestion of large particles, such as some
bacteria ingested by the cell,
major groove. The wide groove on the sur-
face of a DNA double helix created by the
stacking of base pairs and the resulting twist
in the sugar-phosphate backbones.

MALDI. See matrix- assisted laser desorp-
tion ionization.

mass action ratio (Q). The ratio of the con-
centrations of products to the concentrations
of reactants of a reaction,
mass spectrometry. A technique that deter-
mines the mass of a molecule,
matrix. See mitochondrial matrix,
matrix-assisted laser desorption ionization
(MALDI). A technique in mass spectrome-
try where the target molecule is released from
a solid matrix by a laser beam,
maximum velocity ( V max ) • The initial veloc-
ity of a reaction when the enzyme is saturated
with substrate, that is, when all the enzyme is
in the form of an enzyme-substrate complex,
melting curve. A plot of the change in ab-
sorbance versus temperature for a DNA mol-
ecule. The change in absorbance indicates
unfolding of the double helix,
melting point (T m ). The midpoint of the
temperature range in which double-stranded
DNA is converted to single- stranded DNA or
a protein is converted from its native form to
the denatured state.

membrane. A lipid bilayer containing associ-
ated proteins that serves to delineate and com-
partmentalize cells or organelles. Biological
membranes are also the site of many important

GLOSSARY OF BIOCHEMICAL TERMS 759

biochemical processes related to energy trans-
duction and intracellular signaling.

membrane-associated electron transport.

See electron transport.

membrane potential ( A \jj ) . The charge sep-
aration across a membrane that results from
differences in ionic concentrations on the
two sides of the membrane,
messenger ribonucleic acid. See mRNA.
metabolic fuel. A small compound that can
be catabolized to release energy. In multicellu-
lar organisms, metabolic fuels may be trans-
ported between tissues,
metabolically irreversible reaction. A reac-
tion in which the value of the mass action
ratio is two or more orders of magnitude
smaller than the value of the equilibrium
constant. The Gibbs free energy change for
such a reaction is a large negative number;
thus, the reaction is essentially irreversible,
metabolism. The sum total of biochemical
reactions carried out by an organism,
metabolite. An intermediate in the synthesis
or degradation of biopolymers and their
component units.

metabolite channeling. Transfer of the
product of one reaction of a multifunctional
enzyme or a multienzyme complex directly
to the next active site or enzyme without en-
tering the bulk solvent. Channeling increases
the rate of a reaction pathway by decreasing
the transit time for an intermediate to reach
the next enzyme and by producing high local
concentrations of the intermediate,
metalloenzyme. An enzyme that contains
one or more firmly bound metal ions. In
some cases, such metal ions constitute part of
the active site of the enzyme and are active
participants in catalysis,
micelle. An aggregation of amphipathic
molecules in which the hydrophilic portions
of the molecules project into the aqueous
environment and the hydrophobic portions
associated with one another in the interior of
the structure to minimize contact with water
molecules.

Michaelis constant (K m ). The concentra-
tion of substrate that results in an initial
velocity (v 0 ) equal to one-half the maximum
velocity (V^x) for a given reaction.
Michaelis-Menten equation. A rate equation
relating the initial velocity (v 0 ) of an enzy-
matic reaction to the substrate concentration
([S]), the maximum velocity (Vm^), and the
Michaelis constant ( K m ).
microfilament. See actin filament,
microtubule. A protein filament composed
of a and b tubulin heterodimers. Micro-
tubules are components of the cytoskeletal
network and can form structures capable of
directed movement.

minor groove. The narrow groove on the
surface of a DNA double helix created by the
stacking of base pairs and the resulting twist
in the sugar-phosphate backbones.

mismatch repair. Restoration of the normal
nucleotide sequence in a DNA molecule con-
taining mismatched bases. In mismatch re-
pair, the correct strand is recognized, a
portion of the incorrect strand is excised, and
correctly base-paired, double- stranded DNA
is synthesized by the actions of DNA poly-
merase and DNA ligase.
missense mutation. An alteration in DNA
that involves the substitution of one nu-
cleotide for another, resulting in a change in
the amino acid specified by that codon,
mitochondrial matrix. The gel-like phase
enclosed by the inner membrane of the mito-
chondrion. The mitochondrial matrix con-
tains many enzymes involved in aerobic
energy metabolism.

mitochondrion. An organelle that is the
main site of oxidative energy metabolism in
most eukaryotic cells. Mitochondria contain
an outer and an inner membrane, the latter
characteristically folded into cristae.
mixed inhibition. A form of enzyme inhibi-
tion where both K m and V max are affected,
molar mass. The weight in grams of one
mole of a compound,
molecular chaperone. See chaperone,
molecular crowding. The decrease in diffu-
sion rate that occurs when molecules collide
with each other.

molecular weight. See relative molecular
mass.

monocistronic mRNA. An mRNA molecule
that encodes only a single polypeptide. Most
eukaryotic mRNA molecules are mono-
cistronic.

monomer. 1 . A small compound that be-
comes a residue when polymerized with
other monomers. 2. A single subunit of a
multisubunit protein.

monosaccharide. A simple sugar of three or
more carbon atoms with the empirical for-
mula (CH 2 0) n .

monounsaturated fatty acid. An unsaturated
fatty acid with a single carbon- carbon double
bond.

motif. A combination of secondary structure
that appears in a number of different proteins.
Also known as supersecondary structure.

M r . See relative molecular mass.
mRNA. A class of RNA molecules that serve
as templates for protein synthesis.
mRNA precursor. A class of RNA molecules
synthesized by eukaryotic RNA polymerase
II. mRNA precursors are processed posttran-
scriptionally to produce mature messenger
RNA.

mucin. A high-molecular-weight O-linked
glycoprotein containing as much as 80% car-
bohydrate by mass. Mucins are extended,
negatively charged molecules that contribute
to the viscosity of mucus, the fluid found on
the surfaces of the gastrointestinal, genitouri-
nary, and respiratory tracts.

multienzyme complex. An oligomeric pro-
tein that catalyzes several metabolic reactions,
mutagen. An agent that can cause DNA
damage.

mutation. A heritable change in the se-
quence of nucleotides in DNA that causes a
permanent alteration of genetic information,
near-equilibrium reaction. A reaction in
which the value of the mass action ratio is
close to the value of the equilibrium constant.
The Gibbs free energy change for such a reac-
tion is small; thus, the reaction is reversible.
Nernst equation. An equation that relates
the observed change in reduction potential
( A£) to the change in standard reduction po-
tential (A E°') of a reaction,
neutral phospholipids. Glycerophospholipids,
such as phosphatidyl choline, having no net
charge.

neutral solution. An aqueous solution that
has a pH value of 7.0.

nick translation. The process in which DNA
polymerase binds to a gap between the 3' end
of a nascent DNA chain and the 5 ' end of the
next RNA primer, catalyzes hydrolytic re-
moval of ribonucleotides using 5' — » 3' ex-
onuclease activity, and replaces them with
deoxyribonucleotides using 5 ' —> 3 ' poly-
merase activity.

nitrogen cycle. The flow of nitrogen from
N 2 to nitrogen oxides (NOP and NC>P) am-
monia, nitrogenous biomolecules, and back
to N 2 .

nitrogen fixation. The reduction of atmos-
pheric nitrogen to ammonia. Biological ni-
trogen fixation occurs in only a few species of
bacteria and algae.

N-linked oligosaccharide. An oligosaccha-
ride chain attached to a protein through co-
valent bonds to the amide nitrogen atom of
side chain of asparagine residues. The
oligosaccharide chains of N-linked glycopro-
teins contain a core pentasaccharide of two
N-acetylglucosamine residues and three
mannose residues.

NMR spectroscopy. See nuclear magnetic
resonance spectroscopy.

noncompetitive inhibition. Inhibition of an
enzyme-catalyzed reaction by a reversible in-
hibitor that binds to either the enzyme or the
enzyme- substrate complex.

nonessential amino acid. An amino acid
that an animal can produce in sufficient
quantity to meet metabolic needs.

nonhomologous recombination. Recombina-
tion between unrelated sequences that do not
share significant sequence similarity.

nonrepetitive structure. An element of pro-
tein structure in which consecutive residues
do not have a single repeating conformation.

nonsense mutation. An alteration in DNA
that involves the substitution of one nu-
cleotide for another, changing a codon that
specifies an amino acid to a termination

760 GLOSSARY OF BIOCHEMICAL TERMS

codon. A nonsense mutation results in pre-
mature termination of a protein s synthesis.

N-terminus. The amino acid residue bear-
ing a free a:-amino group at one end of a pep-
tide chain. In some proteins, the N-terminus
is blocked by acylation. The N - terminal
residue is usually assigned the residue num-
ber 1. Also known as the amino terminus.

nuclear envelope. The double membrane
that surrounds the nucleus and contains pro-
tein-lined nuclear pore complexes that regu-
late the import and export of material to and
from the nucleus. The outer membrane of
the nuclear envelope is continuous with the
endoplasmic reticulum; the inner membrane
is lined with filamentous proteins, constitut-
ing the nuclear lamina.

nuclear magnetic resonance spectroscopy
(NMR spectroscopy). A technique used to
study the structures of molecules in solution.
In nuclear magnetic resonance spectroscopy,
the absorption of electromagnetic radiation
by molecules in magnetic fields of varying
frequencies is used to determine the spin
states of certain atomic nuclei.

nuclease. An enzyme that catalyzes hydroly-
sis of the phosphodiester linkages of a
polynucleotide chain. Nucleases can be clas-
sified as endonucleases and exonucleases.

nucleic acid. A polymer composed of nu-
cleotide residues linked in a linear sequence
by 3' -5' phosphodiester linkages. DNA and
RNA are nucleic acids composed of deoxyri-
bonucleotide residues and ribonucleotide
residues, respectively.

nucleoid region. The region within a prokary-
otic cell that contains the chromosome.

nucleolus. The region of the eukaryotic nu-
cleus where rRNA transcripts are processed
and ribosomes are assembled.

nucleophile. An electron-rich species that is
negatively charged or contains unshared elec-
tron pairs and is attracted to chemical species
that are positively charged or electron-defi-
cient (electrophiles).

nucleophilic substitution. A reaction in
which one nucleophile (e.g., Y®) displaces
another (e.g.,X®).

nucleoside. A purine or pyrimidine N- gly-
coside of ribose or deoxyribose.
nucleosome. A DNA-protein complex that
forms the fundamental unit of chromatin. A
nucleosome consists of a nucleosome core
particle (approximately 146 base pairs of
DNA plus a histone octamer), linker DNA
(approximately 54 base pairs), and histone
H 1 (which binds the core particle and linker
DNA).

nucleosome core particle. A DNA-protein
complex composed of approximately 146
base pairs of DNA wrapped around an oc-
tamer of histones (two each of H2A, H2B,
H3, and H4).

nucleotide. The phosphate ester of a nucleo-
side, consisting of a nitrogenous base linked

to a pentose phosphate. Nucleotides are the
monomeric units of nucleic acids,
nucleus. An organelle that contains the
principal genetic material of eukaryotic cells
and functions as the major site of RNA syn-
thesis and processing.

obligate aerobe. An organism that requires
the presence of oxygen for survival,
obligate anaerobe. An organism that requires
an oxygen-free environment for survival.
Okazaki fragments. Relatively short strands
of DNA that are produced during discontin-
uous synthesis of the lagging strand of DNA.
oligomer. A multisubunit molecule whose
arrangement of subunits always has a defined
stoichiometry and almost always displays
symmetry.

oligonucleotide. A polymer of several (up to
about 20) nucleotide residues linked by phos-
phodiester bonds.

oligopeptide. A polymer of several (up to
about 20) amino acid residues linked by
peptide bonds.

oligosaccharide. A polymer of 2 to about 20
monosaccharide residues linked by glycosidic
bonds.

oligosaccharide processing. The enzyme-
catalyzed addition and removal of saccharide
residues during the maturation of a glyco-
protein.

O-linked oligosaccharide. An oligosaccha-
ride attached to a protein through a covalent
bond to the hydroxyl oxygen atom of a serine
or threonine residue.

open reading frame. A stretch of nucleotide
triplets that contains no termination codons.
Protein- encoding regions are examples of
open reading frames.

operator. A DNA sequence to which a spe-
cific repressor protein binds, thereby block-
ing transcription of a gene or operon.
operon. A bacterial transcriptional unit
consisting of several different coding regions
cotranscribed from one promoter.

ordered sequential reaction. A reaction in
which both the binding of substrates to an
enzyme and the release of products from the
enzyme follow an obligatory order,
organelle. Any specialized membrane-
bounded structure within a eukaryotic cell.
Organelles are uniquely organized to per-
form specific functions.

origin of replication. A DNA sequence at
which replication is initiated.

osmosis. The movement of solvent mole-
cules from a less concentrated solution to an
adjacent, more concentrated solution,
osmotic pressure. The pressure required to
prevent the flow of solvent from a less con-
centrated solution to a more concentrated
solution.

oxidase. An enzyme that catalyzes an oxida-
tion-reduction reaction in which 0 2 is the
electron acceptor. Oxidases are members of

the IUBMB class of enzymes known as oxi-
doreductases.

oxidation. The loss of electrons from a sub-
stance through transfer to another substance
(the oxidizing agent). Oxidations can take
several forms, including the addition of oxy-
gen to a compound, the removal of hydrogen
from a compound to create a double bond, or
an increase in the valence of a metal ion.
oxidative phosphorylation. See electron
transport.

oxidizing agent. A substance that accepts
electrons in an oxidation- reduction reaction
and thereby becomes reduced,
oxidoreductase. An enzyme that catalyzes an
oxidation -reduction reaction. Some oxidore-
ductases are known as dehydrogenases, oxi-
dases, peroxidases, oxygenases, or reductases,
oxygenation. The reversible binding of oxy-
gen to a macromolecule.

A P- See protonmotive force.

PAGE. See polyacrylamide gel electrophoresis,
passive transport. The process by which a
solute specifically binds to a transport pro-
tein and is transported across a membrane,
moving with the solute concentration gradi-
ent. Passive transport occurs without the ex-
penditure of energy. Also known as facilitated
diffusion.

Pasteur effect. The slowing of glycolysis in
the presence of oxygen,
pathway. A sequence of metabolic reactions,
pause site. A region of a gene where tran-
scription slows. Pausing is exaggerated at
palindromic sequences, where newly synthe-
sized RNA can form a hairpin structure.

PCR. See polymerase chain reaction,
pentose phosphate pathway. A pathway by
which glucose 6-phosphate is metabolized to
generate NADPH and ribose 5 -phosphate. In
the oxidative stage of the pathway, glucose 6-
phosphate is converted to ribulose 5-phosphate
and C0 2 rating two molecules of NADPH. In
the nonoxidative stage, ribulose 5 -phosphate
can be isomerized to ribose 5 -phosphate or
converted to intermediates of glycolysis. Also
known as the hexose monophosphate shunt,
peptide. Two or more amino acids covalently
joined in a linear sequence by peptide bonds,
peptide bond. The covalent secondary
amide linkage that joins the carbonyl group
of one amino acid residue to the amino ni-
trogen of another in peptides and proteins,
peptide group. The nitrogen and carbon
atoms involved in a peptide bond and their
four substituents: the carbonyl oxygen atom,
the amide hydrogen atom, and the two adja-
cent a-carbon atoms.

peptidoglycan. A macromolecule contain-
ing a heteroglycan chain of alternating N-
acetylglucosamine and N-acetylmuramic
acid cross-linked to peptides of varied com-
position. Peptidoglycans are the major com-
ponents of the cell walls of many bacteria,
peptidyl site. See P site.

GLOSSARY OF BIOCHEMICAL TERMS 761

peptidyl transferase. The enzymatic activity
responsible for the formation of a peptide
bond during protein synthesis.
peptidyl-tRNA. The tRNA molecule to
which the growing peptide chain is attached
during protein synthesis,
peripheral membrane protein. A membrane
protein that is weakly bound to the interior or
exterior surface of a membrane through ionic
interactions and hydrogen bonding with the
polar heads of the membrane lipids or with an
integral membrane protein. Also known as an
extrinsic membrane protein,
periplasmic space. The region between the
plasma membrane and the cell wall in bacteria,
permeability coefficient. A measure of the
ability of an ion or small molecule to diffuse
across a lipid bilayer.

peroxisome. An organelle in all animal and
many plant cells that carries out oxidation re-
actions, some of which produce the toxic
compound hydrogen peroxide (H 2 0 2 ). Per-
oxisomes contain the enzyme catalase, which
catalyzes the breakdown of toxic H 2 0 2 to
water and 0 2 .

pH. A logarithmic quantity that indicates
the acidity of a solution, that is, the concen-
tration of hydronium ions in solution. pH is
defined as the negative logarithm of the hy-
dronium ion concentration.
pH optimum. In an enzyme-catalyzed re-
action, the pH at the point of maximum
catalytic activity,
phage. See bacteriophage,
phase-transition temperature (T m ). The
midpoint of the temperature range in which
lipids or other macromolecular aggregates
are converted from a highly ordered phase or
state (such as a gel) to a less- ordered state
(such as a liquid crystal).
c f> (phi). The angle of rotation around the
bond between the a - carbon and the nitrogen
of a peptide group.

phosphagen. A “high energy” phosphate
storage molecule found in animal muscle
cells. Phosphagens are phosphoamides and
have a higher phosphoryl-group-transfer
potential than ATP.

phosphatase. An enzyme that catalyzes
the hydrolytic removal of a phosphoryl
group.

phosphatidate. A glycerophospholipid that
consists of two fatty acyl groups esterified to
C-l and C-2 of glycerol 3-phosphate. Phos-
phatidates are metabolic intermediates in the
biosynthesis or breakdown of more complex
glycerophospholipids.

phosphoanhydride. A compound formed
by condensation of two phosphate groups,
phosphodiester linkage. A linkage in nucleic
acids and other molecules in which two alco-
holic hydroxyl groups are joined through a
phosphate group.

phosphoester linkage. The bond by which a
phosphoryl group is attached to an alcoholic
or phenolic oxygen.

phospholipid. A lipid containing a phos-
phate moiety.

phosphorolysis. Cleavage of a bond within
a molecule by group transfer to an oxygen
atom of phosphate.

phosphorylase. An enzyme that catalyzes
the cleavage of its substrate(s) via nucle-
ophilic attack by inorganic phosphate (Pj)
(i.e., via phosphorolysis).
phosphorylation. A reaction involving the
addition of a phosphoryl group to a molecule,
phosphoryl group transfer potential. A
measure of the ability of a compound to
transfer a phosphoryl group to another com-
pound. Under standard conditions, group
transfer potentials have the same values as
the standard free energies of hydrolysis but
are opposite in sign.

photoautotroph. A photosynthetic organ-
ism that can utilize C0 2 as its main carbon
source.

photon. A quantum of light energy,
photophosphorylation. The light-dependent
formation of ATP from ADP and Pj catalyzed
by chloroplast ATP synthase,
photoheterotroph. Photosynthetic organ-
ism that requires organic molecules as a car-
bon source.

photoreactivation. The direct repair of
damaged DNA by an enzyme that is activated
by visible light.

photorespiration. The light-dependent up-
take of 0 2 and the subsequent metabolism of
phosphoglycolate that occurs primarily in C 3
photosynthetic plants. Photorespiration can
occur because 0 2 competes with C0 2 for
the active site of ribulose 1,5-fo'sphosphate
carboxylase-oxygenase, the enzyme that cat-
alyzes the first step of the reductive pentose
phosphate cycle.

photosynthesis. The conversion of light en-
ergy (photons) to chemical energy in the
form of ATP and/or NADPH.
photosystem. A functional unit of the light-
dependent electron-transfer reactions of
photosynthesis. Each membrane-embedded
photosystem contains a reaction center, which
forms the core of the photosystem, and a pool
of light- absorbing antenna pigments,
phototroph. An organism that can convert
light energy into chemical potential energy
(i.e., an organism capable of photosynthesis),
physiological pH. The normal pH of human
blood, which is 7.4.
pi. See isoelectric point,
ping-pong reaction. A reaction in which an
enzyme binds one substrate and releases a
product, leaving a substituted enzyme that
then binds a second substrate and releases a
second product, thereby restoring the en-
zyme to its original form,
pitch. The axial distance for one complete
turn of a helical structure,
p K a . A logarithmic value that indicates the
strength of an acid, p is defined as the

negative logarithm of the acid dissociation
constant, K a .

plasma membrane. The membrane that
surrounds the cytoplasm of a cell and thus
defines the perimeter of the cell,
plasmalogen. A glycerophospholipid that
has a hydrocarbon chain linked to C-l of
glycerol 3 -phosphate through a vinyl ether
linkage. Plasmalogens are found in the cen-
tral nervous system and in peripheral nerve
and muscle tissue.

plasmid. A relatively small, extrachromo-
somal DNA molecule that is capable of au-
tonomous replication. Plasmids are usually
closed, circular, double-stranded DNA
molecules.

P:0 ratio. The ratio of molecules of ADP
phosphorylated to atoms of oxygen reduced
during oxidative phosphorylation,
polar. Having uneven distribution of charge.
A molecule or functional group is polar if its
center of negative charge does not coincide
with its center of positive charge,
poly A tail. A stretch of polyadenylate, up to
250 nucleotide residues long, that is added to
the 3' end of a eukaryotic mRNA molecule
following transcription,
polyacrylamide gel electrophoresis (PAGE).
A technique used to separate molecules of
different net charge and/or size based on
their migration through a highly cross-linked
gel matrix in an electric field,
polycistronic mRNA. An mRNA molecule
that contains multiple coding regions. Many
prokaryotic mRNA molecules are polycistronic.
polymerase chain reaction (PCR). A
method for amplifying the amount of DNA in
a sample and for enriching a particular DNA
sequence in a population of DNA molecules.
In the polymerase chain reaction, oligonu-
cleotides complementary to the ends of the
desired DNA sequence are used as primers for
multiple rounds of DNA synthesis,
polynucleotide. A polymer of many (usually
more than 20) nucleotide residues linked by
phosphodiester bonds.

polypeptide. A polymer of many (usually
more than 20) amino acid residues linked by
peptide bonds.

polyribosome. See polysome,
polysaccharide. A polymer of many (usually
more than 20) monosaccharide residues
linked by glycosidic bonds. Polysaccharide
chains can be linear or branched,
polysome. The structure formed by the
binding of many translation complexes to a
large mRNA molecule. Also known as a
polyribosome.

polyunsaturated fatty acid. An unsaturated
fatty acid with two or more carbon-carbon
double bonds,
pore. See channel.

posttranscriptional processing. RNA pro-
cessing that occurs after transcription is
complete.

762 GLOSSARY OF BIOCHEMICAL TERMS

posttranslational modification. Covalent
modification of a protein that occurs after
synthesis of the polypeptide is complete,
prenylated protein. A lipid-anchored pro-
tein that is covalently linked to an isoprenoid
moiety via the sulfur atom of a cysteine
residue at the C-terminus of the protein,
primary structure. The sequence in which
residues are covalently linked to form a poly-
meric chain.

primary transcript. A newly synthesized
RNA molecule before processing,
primase. An enzyme in the primosome that
catalyzes the synthesis of short pieces of RNA
about 10 residues long. These oligonu-
cleotides are the primers for synthesis of
Okazaki fragments.

primosome. A multiprotein complex, in-
cluding primase and helicase in E. coli , that
catalyzes the synthesis of the short RNA
primers needed for discontinuous DNA syn-
thesis of the lagging strand,
processive enzyme. An enzyme that re-
mains bound to its growing polymeric
product through many polymerization steps
(cf., distributive enzyme),
prochiral atom. An atom with multiple sub-
stituents, two of which are identical. A
prochiral atom can become chiral when one
of the identical substituents is replaced,
prokaryote. An organism, usually a single
cell, which contains no nucleus or internal
membranes (cf., eukaryote),
promoter. The region of DNA where RNA
polymerase binds during transcription
initiation.

prostaglandin. An eicosanoid that has a cy-
clopentane ring. Prostaglandins are meta-
bolic regulators that act in the immediate
neighborhood of the cells in which they are
produced.

prosthetic group. A coenzyme that is tightly
bound to an enzyme. A prosthetic group, un-
like a cosubstrate, remains bound to a spe-
cific site of the enzyme throughout the
catalytic cycle of the enzyme,
protease. An enzyme that catalyzes hydroly-
sis of peptide bonds. The physiological sub-
strates of proteases are proteins,
protein. A biopolymer consisting of one or
more polypeptide chains. The biological
function of each protein molecule depends
not only on the sequence of covalently linked
amino acid residues, but also on its three-
dimensional structure (conformation),
protein coenzyme. A protein that does not
itself catalyze reactions but is required for the
action of certain enzymes,
protein glycosylation. The covalent addition
of carbohydrate to proteins. In N- glycosyla-
tion, the carbohydrate is attached to the
amide group of the side chain of an as-
paragine residue. In O-glycosylation, the car-
bohydrate is attached to the hydroxyl group of
the side chain of a serine or threonine residue.

protein kinase. See kinase,
protein phosphatase. See phosphatase,
proteoglycan. A complex of protein with
glycosaminoglycan chains covalently bound
through their anomeric carbon atoms. Up to
95% of the mass of a proteoglycan may be
glycosaminoglycan.

proteomics. The study of all proteins pro-
duced in a certain cell type, tissue, organ, or
organism.

protonmotive force (A p). The energy
stored in a proton concentration gradient
across a membrane.

proximity effect. The increase in the rate of
a nonenzymatic or enzymatic reaction attrib-
utable to high effective concentrations of re-
actants, which result in more frequent
formation of transition states,
pseudo first-order reaction. A multi-reactant
reaction carried out under conditions where
the rate depends on the concentration of only
one reactant.

pseudogene. A nonexpressed sequence of
DNA that evolved from a protein-encoding
gene. Pseudogenes often contain mutations
in their coding regions and cannot produce
functional proteins.

if/ (psi). The angle of rotation around the
bond between the a-carbon and the carbonyl
carbon of a peptide group.

A ifj. See membrane potential.

P site. Peptidyl site. The site on a ribosome
that is occupied during protein synthesis by a
tRNA molecule attached to the growing
polypeptide chain (peptidyl tRNA).
purine. A nitrogenous base having a two-
ring structure in which a pyrimidine is fused
to imidazole. Adenine and guanine are substi-
tuted purines found in both DNA and RNA.
pyranose. A monosaccharide structure that
forms a six-membered ring as a result of in-
tramolecular hemiacetal formation,
pyrimidine. A nitrogenous base having a
heterocyclic ring that consists of four carbon
atoms and two nitrogen atoms. Cytosine,
thymine, and uracil are substituted pyrim-
idines found in nucleic acids (cytosine in
DNA and RNA, uracil in RNA, and thymine
principally in DNA).

Q. See mass action ratio.

Q cycle. A cyclic pathway proposed to explain
the sequence of electron transfers and proton
movements within Complex III of mitochon-
dria or the cytochrome bf complex in chloro-
plasts. The net result of the two steps of the
Q cycle is oxidation of two molecules of QH 2
or plastoquinol (PQH 2 ); formation of one
molecule of QH 2 or PQH 2 ; transfer of two
electrons; and net translocation of four pro-
tons across the inner mitochondrial mem-
brane to the intermembrane space or across
the thylakoid membrane to the lumen,
quaternary structure. The organization of
two or more polypeptide chains within a
multisubunit protein.

R state. The more active conformation of an
allosteric protein; opposite of T state.
Ramachandran plot. A plot of c versus f val-
ues for amino acid residues in a polypeptide
chain. Certain f and c values are characteris-
tic of different conformations,
random sequential reaction. A reaction in
which neither the binding of substrates to an
enzyme nor the release of products from the
enzyme follows an obligatory order,
rate acceleration. The ratio of the rate con-
stant for a reaction in the presence of enzyme
(fccat) divided by the rate constant for that re-
action in the absence of enzyme (k n ). The
rate acceleration value is a measure of the ef-
ficiency of an enzyme.

rate equation. An expression of the observed
relationship between the velocity of a reaction
and the concentration of each reactant,
rate determining step. The slowest step in a
chemical reaction. The rate determining step
has the highest activation energy among the
steps leading to formation of a product from
the substrate.

reaction center. A complex of proteins, elec-
tron transport cofactors, and a special pair
of chlorophyll molecules that forms the
core of a photosystem. The reaction center
is the site of conversion of photochemical
energy to electrochemical energy during
photosynthesis.

reaction mechanism. The step-by-step
atomic or molecular events that occur during
chemical reactions,
reaction order. See kinetic order,
reaction specificity. The lack of formation
of wasteful by-products by an enzyme. Reac-
tion specificity results in essentially 100%
product yields.

reactive center. The part of a coenzyme to
which mobile metabolic groups are attached,
reading frame. The sequence of nonoverlap-
ping codons of an mRNA molecule that spec-
ifies the amino acid sequence. The reading
frame of an mRNA molecule is determined by
the position where translation begins; usually
an AUG codon.

receptor. A protein that binds a specific
ligand, such as a hormone, leading to some
cellular response.

recombinant DNA. A DNA molecule that
includes DNA from different sources,
recombination. See genetic recombination,
reducing agent. A substance that loses elec-
trons in an oxidation-reduction reaction and
thereby becomes oxidized,
reducing end. The residue containing a free
anomeric carbon in a polysaccharide. A poly-
saccharide usually contains no more than
one reducing end.

reduction. The gain of electrons by a sub-
stance through transfer from another sub-
stance (the reducing agent). Reductions can
take several forms, including the loss of oxy-
gen from a compound, the addition of

GLOSSARY OF BIOCHEMICAL TERMS 763

hydrogen to a double bond of a compound,
or a decrease in the valence of a metal ion.
reduction potential ( E ). A measure of the
tendency of a substance to reduce other sub-
stances. The more negative the reduction po-
tential, the greater the tendency to donate
electrons.

regulated enzyme. An enzyme located at a
critical point within one or more metabolic
pathways, whose activity may be increased or
decreased based on metabolic demand. Most
regulated enzymes are oligomeric,
regulatory protein. A protein that is in-
volved in the regulation of gene expression,
usually at the point of transcription initia-
tion. Repressors and activators are examples
of regulatory proteins.

regulatory site. A ligand-binding site in a
regulatory enzyme distinct from the active
site. Allosteric modulators alter enzyme ac-
tivity by binding to the regulatory site. Also
known as an allosteric site,
relative molecular mass (M r ). The mass of a
molecule relative to l/12th the mass of 12 C.
There are no units associated with the values
for relative molecular mass,
release factor. A protein involved in termi-
nating protein synthesis,
renaturation. The restoration of the native
conformation of a biological macromolecule,
usually resulting in restoration of biological
activity.

replication. The duplication of double-
stranded DNA, during which parental strands
separate and serve as templates for synthesis
of new strands. Replication is carried out by
DNA polymerase and associated factors,
replication fork. The Y-shaped junction
where double-stranded, template DNA is
unwound and new DNA strands are synthe-
sized during replication,
replisome. A multiprotein complex that in-
cludes DNA polymerase, primase, helicase,
single-strand binding protein, and additional
components. The replisomes, located at each
of the replication forks, carry out the poly-
merization reactions of bacterial chromoso-
mal DNA replication.

repressor. A regulatory DNA-binding pro-
tein that prevents transcription by RNA
polymerase.

residue. A single component within a poly-
mer. The chemical formula of a residue is
that of the corresponding monomer minus
the elements of water.

resonance energy transfer. A form of exci-
tation energy transfer between molecules
that does not involve transfer of an electron,
respiratory electron transport chain. A
series of enzyme complexes and associated
cofactors that are electron carriers, passing
electrons from reduced coenzymes or
substrates to molecular oxygen (0 2 )> the
terminal electron acceptor of aerobic
metabolism.

restriction endonuclease. An endonuclease
that catalyzes the hydrolysis of double- strand-
ed DNA at a specific nucleotide sequence.
Type I restriction endonucleases catalyze both
the methylation of host DNA and the cleavage
of nonmethylated DNA, whereas type II re-
striction endonucleases catalyze only the
cleavage of nonmethylated DNA.
restriction map. A diagram showing the size
and arrangement of fragments produced
from a DNA molecule by the action of vari-
ous restriction endonucleases,
reverse transcriptase. A type of DNA poly-
merase that catalyzes the synthesis of a strand
of DNA from an RNA template,
reverse turn. See turn,
ribonucleic acid (RNA). A polymer consist-
ing of ribonucleotide residues joined by
3' -5' phosphodiester bonds. The sugar moi-
ety in RNA is ribose. Genetic information
contained in DNA is transcribed in the syn-
thesis of RNA, some of which (mRNA) is
translated in the synthesis of protein,
ribonucleoprotein. A complex containing
both ribonucleic acid and protein,
ribosome. A large ribonucleoprotein com-
plex composed of multiple ribosomal RNA
molecules and proteins. Ribosomes are the
site of protein synthesis,
ribozyme. An RNA molecule with enzymatic
activity.

rise. The distance between one residue and
the next along the axis of a helical macro -
molecule.

RNA processing. The reactions that trans-
form a primary RNA transcript into a ma-
ture RNA molecule. The three general types
of RNA processing include the removal of
RNA nucleotides from primary transcripts,
the addition of RNA nucleotides not encod-
ed by the gene, and the covalent modifica-
tion of bases.

rRNA. See ribosomal ribonucleic acid.

S. See Svedberg unit.

S. Sec entropy.

salt bridge. See charge- charge interactions,
salvage pathway. A pathway in which a
major metabolite, such as a purine or pyrim-
idine nucleotide, can be synthesized from a
preformed molecular entity, such as a purine
or pyrimidine.

saturated fatty acid. A fatty acid that does
not contain a carbon-carbon double bond.
Schiff base. A complex formed by the re-
versible condensation of a primary amine
with an aldehyde (to form an aldimine) or a
ketone (to form a ketimine).

SDS-PAGE. See sodium dodecyl sulfate-poly-
acrylamide gel electrophoresis,
second messenger. A compound that acts
intracellularly in response to an extracellular
signal.

secondary structure. The regularities in
local conformations within macromolecules.

In proteins, secondary structure is main-
tained by hydrogen bonds between carbonyl
and amide groups of the backbone. In nucle-
ic acids, secondary structure is maintained by
hydrogen bonds and stacking interactions
between the bases.

second-order reaction. A reaction whose
rate depends on the concentrations of two re-
actants.

self-splicing intron. An intron that is ex-
cised in a reaction mediated by the RNA pre-
cursor itself.

sense strand. In double-stranded DNA the
sense strand is the strand that contains
codons. Also called the coding strand. The
opposite strand is called the antisense strand
or the template strand,
sequential reaction. An enzymatic reac-
tion in which all the substrates must be
bound to the enzyme before any product is
released.

sequential theory of cooperativity and al-
losteric regulation. A model of the coopera-
tive binding of identical ligands to oligomeric
proteins. According to the simplest form of
the sequential theory, the binding of a ligand
may induce a change in the tertiary structure
of the subunit to which it binds and may
alter the conformations of neighboring sub-
units to varying extents. Only one subunit
conformation has a high affinity for the
ligand. Also known as the ligand-induced
theory.

Shine-Dalgarno sequence. A purine-rich
region just upstream of the initiation codon
in prokaryotic mRNA molecules. The Shine-
Dalgarno sequence binds to a pyrimidine-
rich sequence in the ribosomal RNA, thereby
positioning the ribosome at the initiation
codon.

i t factor. See a subunit.

c t subunit (sigma subunit). A subunit of
prokaryotic RNA polymerase, which acts as a
transcription initiation factor by binding to
the promoter. Different a subunits are spe-
cific for different promoters. Also known as a
a factor.

signal peptidase. An integral membrane
protein of the endoplasmic reticulum that
catalyzes cleavage of the signal peptide of
proteins translocated to the lumen.

signal peptide. The N-terminal sequence of
residues in a newly synthesized polypeptide
that targets the protein for translocation
across a membrane.

signal transduction. The process whereby
an extracellular signal is converted to an in-
tracellular signal by the action of a mem-
brane-associated receptor, a transducer, and
an effector enzyme.

signal recognition particle (SRP). A eu-
karyotic protein-RNA complex that binds a
newly synthesized peptide as it is extruded
from the ribosome. The signal-recognition
particle is involved in anchoring the ribo-
some to the cytosolic face of the endoplasmic

764 GLOSSARY OF BIOCHEMICAL TERMS

reticulum so that protein translocation to the
lumen can occur.

single-strand binding protein (SSB). A pro-
tein that binds tightly to single-stranded
DNA, preventing the DNA from folding back
on itself to form double-stranded regions.

site-directed mutagenesis. An in vitro pro-
cedure by which one particular nucleotide
residue in a gene is replaced by another, re-
sulting in production of an altered protein
sequence.

site-specific recombination. An example of
recombination that occurs at specific sites in
the genome.

small nuclear ribonucleoprotein (snRNP).

An RNA-protein complex composed of one
or two specific snRNA molecules plus a num-
ber of proteins. snRNPs are involved in splic-
ing mRNA precursors and in other cellular
events.

small RNA. A class of RNA molecules. Some
small RNA molecules have catalytic activity.
Some small nuclear RNA molecules (snRNA)
are components of small nuclear ribonucleo-
proteins (snRNPs).
snRNA. See small nuclear RNA.
snRNP. See small nuclear ribonucleoprotein.
sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE). Polyacrylamide
gel electrophoresis performed in the presence
of the detergent sodium dodecyl sulfate. SDS-
PAGE allows separation of proteins on the
basis of size only rather than charge and size,
solvation. A state in which a molecule or ion
is surrounded by solvent molecules,
solvation sphere. The shell of solvent mole-
cules that surrounds an ion or solute,
special pair. A specialized pair of chloro-
phyll molecules in reaction centers that is the
primary electron donor during the light-de-
pendent reactions of photosynthesis.

specific heat. The amount of heat required
to raise the temperature of 1 gram of a sub-
stance by 1°C.

specificity constant. Seek cat /K m .

sphingolipid. An amphipathic lipid with a
sphingosine (trans-4-sphingenine) back-
bone. Sphingolipids, which include sphin-
gomyelins, cerebrosides, and gangliosides, are
present in plant and animal membranes and
are particularly abundant in the tissues of the
central nervous system.

sphingomyelin. A sphingolipid that consists
of phosphocholine attached to the C-l hy-
droxyl group of a ceramide. Sphingomyelins
are present in the plasma membranes of most
mammalian cells and are a major component
of myelin sheaths.

splice site. The conserved nucleotide se-
quence surrounding an exon-intron junction.
It includes the site where the RNA molecule is
cleaved during intron excision.

spliceosome. The large protein-RNA com-
plex that catalyzes the removal of introns from

mRNA precursors. The spliceosome is com-
posed of small nuclear ribonucleoproteins.
splicing. The process of removing introns
and joining exons to form a continuous RNA
molecule.

SRR See signal recognition particle.

SSB. See single-strand binding protein,
stacking interactions. The weak noncovalent
forces between adjacent bases or base pairs in
single- stranded or double- stranded nucleic
acids, respectively. Stacking interactions con-
tribute to the helical shape of nucleic acids,
standard Gibbs free energy change (AG°')«
The free energy change for a reaction under
biochemical standard state conditions,
standard reduction potential (E°')« A
measure of the tendency of a substance to re-
duce other substances under biochemical
standard state conditions,
standard state. A set of reference conditions
for a chemical reaction. In biochemistry, the
standard state is defined as a temperature of
298 K (25°C), a pressure of 1 atmosphere, a
solute concentration of 1.0 M, and a pH of 7.0.
starch. A homopolymer of glucose residues
that is a storage polysaccharide in plants.
There are two forms of starch: amylose, an
unbranched polymer of glucose residues
joined by cr-(1^4) linkages; and amy-
lopectin, a branched polymer of glucose
residues joined by a-( 1 — » 4) linkages with
a-( 1 — > 6) linkages at branch points,
steady state. A state in which the rate of syn-
thesis of a compound is equal to its rate of
utilization or degradation,
stem-loop. See hairpin,
stereoisomers. Compounds with the same
molecular formula but different spatial
arrangements of their atoms,
stereospecificity. The ability of an enzyme
to recognize and act upon only a single
stereoisomer of a substrate,
steroid. A lipid containing a fused, four-ring
isoprenoid structure.

sterol. A steroid containing a hydroxyl group,
stomata. Structures on the surface of a leaf
through which carbon dioxide diffuses di-
rectly into photosynthetic cells,
stop codon. See termination codon,
strand invasion. The exchange of single
strands of DNA from two nicked molecules
having homologous nucleotide sequences,
stroma. The interior of a chloroplast corre-
sponding to the cytoplasm of the ancestral
cyanobacterium.

stromal lamellae. Regions of the thylakoid
membrane that are in contact with the stroma,
substrate. A reactant in a chemical reaction.
In enzymatic reactions, substrates are specifi-
cally acted upon by enzymes, which catalyze
the conversion of substrates to products,
substrate cycle. A pair of opposing reactions
that catalyzes a cycle between two pathway
intermediates.

substrate level phosphorylation. Phosphory-
lation of a nucleoside diphosphate by
transfer of a phosphoryl group from a non-
nucleotide substrate.

supercoil. A topological arrangement assumed
by over- or underwound double-stranded DNA.
Underwinding gives rise to negative supercoils;
overwinding produces positive supercoils,
supersecondary structure. See motif.
Svedberg unit (S). A unit of 10 -13 second
used for expressing the sedimentation coeffi-
cient, a measure of the rate at which a large
molecule or particle sediments in an ultra-
centrifuge. Large S values usually indicate
large masses.

symport. The cotransport of two different
species of ions or molecules in the same direc-
tion across a membrane by a transport protein,
synonymous codons. Different codons that
specify the same amino acid,
synthase. A common name for an enzyme,
often a transferase, that catalyzes a synthetic
reaction.

synthetase. An enzyme that catalyzes the join-
ing of two substrates and requires the input of
the chemical potential energy of a nucleoside
triphosphate. Synthetases are members of the
IUBMB class of enzymes known as ligases.

T state. The less active conformation of an
allosteric protein; opposite of R state.

TATA box. An A/T-rich DNA sequence
found within the promoter of both prokary-
otic and eukaryotic genes,
template strand. The strand of DNA within
a gene whose nucleotide sequence is comple-
mentary to that of the transcribed RNA. Dur-
ing transcription, RNA polymerase binds to
and moves along the template strand in the
3' — » 5' direction, catalyzing the synthesis of
RNA in the 5' —> 3' direction,
termination codon. A codon that is recog-
nized by specific proteins that cause newly
synthesized peptides to be released from the
translation machinery thus terminating
translation. The three termination codons
(UAG, UAA, and UGA) are also known as
stop codons.

termination sequence. A sequence at the 3'
end of a gene that mediates transcription
termination.

tertiary structure. The compacting of poly-
meric chains into one or more domains with-
in a macromolecule. In proteins, tertiary
structure is stabilized mainly by hydrophobic
interactions between side chains,
thermodynamics. The branch of physical
science that studies transformations of heat
and energy.

30 nm fiber. A chromatin structure in which
nucleosomes are coiled into a solenoid 30 nm
in diameter.

— 35 region. A sequence found within the
promoter of some prokaryotic genes about
30 to 35 base pairs upstream of the transcrip-
tion initiation site.

GLOSSARY OF BIOCHEMICAL TERMS 765

3 10 helix. A secondary structure of proteins,
consisting of a helix in which the carbonyl
oxygen of each amino acid residue (residue
n) forms a hydrogen bond with the amide hy-
drogen of the third residue further toward
the C-terminus of the polypeptide chain
(residue n + 3).

thylakoid lamella. See thylakoid membrane,
thylakoid membrane. A highly folded, con-
tinuous membrane network suspended in
the aqueous matrix of the chloroplast. The
thylakoid membrane is the site of the light-
dependent reactions of photosynthesis,
which lead to the formation of NADPH and
ATP. Also known as the thylakoid lamella.

T m . See melting point and phase-transition
temperature.

topoisomerase. An enzyme that alters the
supercoiling of a DNA molecule by cleaving a
phosphodiester linkage in either one or both
strands, rewinding the DNA, and resealing
the break. Some topoisomerases are also
known as DNA gyrases.
topology. 1 . The arrangement of membrane-
spanning segments and connecting loops in
an integral membrane protein. 2. The overall
morphology of a nucleic acid molecule.

Ti/fC arm. The stem-and-loop structure in a
tRNA molecule that contains the sequence
ribothymidylate-pseudouridylate-cytidylate
W C).

trace element. An element required in very
small quantities by living organisms. Exam-
ples include copper, iron, and zinc,
transaminase. An enzyme that catalyzes the
transfer of an amino group from an a - amino
acid to an a-keto acid. Transaminases require
the coenzyme pyridoxal phosphate. They are
also called aminotransferases,
transcription. The copying of biological in-
formation from a double- stranded DNA
molecule to a single-stranded RNA molecule,
catalyzed by a transcription complex consist-
ing of RNA polymerase and associated
factors.

transcription bubble. A short region of
double- stranded DNA that is unwound by
RNA polymerase during transcription,
transcription factor. A protein that binds to
the promoter region, to RNA polymerase, or
to both during assembly of the transcription
initiation complex. Some transcription
factors remain bound during RNA chain
elongation.

transcription initiation complex. The com-
plex of RNA polymerase and other factors
that assembles at the promoter at the start of
transcription.

transcriptional activator. A regulatory DNA-
binding protein that enhances the rate of
transcription by increasing the activity of
RNA polymerase at specific promoters,
transducer. The component of a signal-
transduction pathway that couples receptor-
ligand binding with generation of a second
messenger catalyzed by an effector enzyme.

transfer ribonucleic acid. See tRNA.
transferase. An enzyme that catalyzes a
group -transfer reaction. Transferases often
require a coenzyme.

transition state. An unstable, high-energy
arrangement of atoms in which chemical
bonds are being formed or broken. Transi-
tion states have structures between those of
the substrates and the products of a reaction,
transition-state analog. A compound that
resembles a transition state. Transition-state
analogs characteristically bind extremely
tightly to the active sites of appropriate en-
zymes and thus act as potent inhibitors,
transition-state stabilization. The increased
binding of transition states to enzymes relative
to the binding of substrates or products. Tran-
sition-state stabilization lowers the activation
energy and thus contributes to catalysis,
translation. The synthesis of a polypeptide
whose sequence reflects the nucleotide se-
quence of an mRNA molecule. Amino acids
are donated by activated tRNA molecules,
and peptide bond synthesis is catalyzed by
the translation complex, which includes the
ribosome and other factors,
translation complex. The complex of a ri-
bosome and protein factors that carries out
the translation of mRNA in vivo,
translation initiation complex. The complex
of ribosomal subunits, an mRNA template, an
initiator tRNA molecule, and initiation factors
that assembles at the start of protein synthesis,
translation initiation factor. A protein in-
volved in the formation of the initiation
complex at the start of protein synthesis,
translocation. 1 . The movement of the ri-
bosome by one codon along an mRNA mole-
cule. 2. The movement of a polypeptide
through a membrane.

transposon. A mobile genetic element that
jumps between chromosomes or parts of a
chromosome by taking advantage of recom-
bination mechanisms. Also known as a trans-
posable element.

transverse diffusion. The passage of lipid or
protein molecules from one leaflet of a lipid
bilayer to the other leaflet. Unlike lateral dif-
fusion within one leaflet of a bilayer, trans-
verse diffusion is extremely slow,
triacylglycerol. A lipid containing three
fatty acyl residues esterified to glycerol. Fats
and oils are mixtures of triacylglycerols. For-
merly known as a triglyceride,
tricarboxylic acid cycle. See citric acid cycle,
triglyceride. See triacylglycerol.
triose. A three-carbon sugar.
tRNA. A class of RNA molecules that carry
activated amino acids to the site of protein
synthesis for incorporation into growing
peptide chains. tRNA molecules contain an
anticodon that recognizes a complementary
codon in mRNA.

turn (in proteins). A protein loop of 4-5
residues that causes a change in the direction
of a polypeptide chain in a folded protein.

turnover. The dynamic metabolic steady
state in which molecules are degraded and re-
placed by newly synthesized molecules,
turnover number. See catalytic constant,
twist. The angle of rotation between adja-
cent residues within a helical macromolecule,
type I reaction center. The special pair of
chlorophyll molecules and associated elec-
tron transfer chain found in photosystem I.
type II reaction center. The reaction center
found in photosystem II.
uncompetitive inhibition. Inhibition of an
enzyme-catalyzed reaction by a reversible in-
hibitor that binds only to the enzyme- sub-
strate complex, not to the free enzyme,
uncouplers. See uncoupling agent,
uncoupling agent. A compound that dis-
rupts the usual tight coupling between elec-
tron transport and phosphorylation of ADR
uniport. The transport of a single type of
solute across a membrane by a transport
protein.

unsaturated fatty acid. A fatty acid with at
least one carbon-carbon double bond. An un-
saturated fatty acid with only one carbon-car-
bon double bond is called a monounsaturated
fatty acid. A fatty acid with two or more car-
bon-carbon double bonds is called a polyun-
saturated fatty acid. In general, the double
bonds of unsaturated fatty acids are of the cis
configuration and are separated from each
other by methylene ( — CH 2 — ) groups,
urea cycle. A metabolic cycle consisting of
four enzyme-catalyzed reactions that con-
verts nitrogen from ammonia and aspartate
to urea. Four ATP equivalents are consumed
during formation of one molecule of urea,
v. Sec velocity.
v 0 . See initial velocity.

vacuole. A fluid-filled organelle in plant
cells that is a storage site for water, ions, or
nutrients.

van der Waals force. A weak intermolecular
force produced between neutral atoms by
transient electrostatic interactions. Van der
Waals attraction is strongest when atoms are
separated by the sum of their van der Waals
radii; strong van der Waals repulsion pre-
cludes closer approach,
van der Waals radius. The effective size of
an atom. The distance between the nuclei of
two nonbonded atoms at the point of maxi-
mal attraction is the sum of their van der
Waals radii.

variable arm. The arm of a tRNA molecule
that is located between the anticodon arm
and the TiffC arm. The variable arm can range
in length from about 3 to 21 nucleotides,
velocity (V). The rate of a chemical reaction,
expressed as amount of product formed per
unit time.

very low density lipoprotein (VLDL). A

type of plasma lipoprotein that transports
endogenous triacylglycerols, cholesterol, and
cholesteryl esters from the liver to the tissues.

766 GLOSSARY OF BIOCHEMICAL TERMS

vitamin. An organic micronutrient that
cannot be synthesized by an animal and must
be obtained in the diet. Many coenzymes are
derived from vitamins.

VLDL. See very low density lipoprotein.
V nvdX . See maximum velocity,
wax. A nonpolar ester that consists of a long
chain monohydroxylic alcohol and a long
chain fatty acid.

wobble position. The 5' position of an anti-
codon, where non-Watson-Crick base pair-
ing with a nucleotide in mRNA is permitted.

The wobble position makes it possible for a
tRNA molecule to recognize more than one
codon.

X-ray crystallography. A technique used to
determine secondary, tertiary, and quater-
nary structures of biological macromole-
cules. In X-ray crystallography, a crystal of
the macromolecule is bombarded with X
rays, which are diffracted and then detected
electronically or on a film. The atomic struc-
ture is deduced by mathematical analysis of
the diffraction pattern.

Z-DNA. A conformation of oligonucleotide
sequences containing alternating deoxycytidy-
late and deoxyguanylate residues. Z-DNA is a
left-handed double helix containing approxi-
mately 12 base pairs per turn,
zero-order reaction. A reaction whose rate
is independent of reactant concentration.
Z-scheme. A zigzag scheme that illustrates the
reduction potentials associated with electron
flow through photosynthetic electron carriers,
zwitterion. A molecule containing negatively
and positively charged groups.

Photo and Illustration Credits

Chapter 1 Page 2 top, Science Photo Library/Photo Researchers, Inc.; 2 middle,
Photos 12/Alamy; 2 bottom, Science Photo Library/Photo Researchers, Inc.;

3 top, Corbis; 3 bottom, Shutterstock; 11, Shutterstock; 12, Manuscripts 8c
Archives — Yale University Library; 15 top, SSPL/The Image Works; 15 bottom,
Richard Bizley/Photo Researchers, Inc.; 18 top, Lee D. Simon/Photo Researchers,
Inc.; 18 bottom, National Library of Medicine Profiles in Science; 20, Matthew
Daniels, Wellcome Images; 22, Dr. Torsten Wittmann/Photo Researchers, Inc.;
and 23, David S. Goodsell, the RCSB Protein Data Bank. Coordinates from
PDB entry latn.

Chapter 2 Page 28 top, NASA; 28 bottom, Michael Charters; 31, iStockphoto;
32, NOAA; 33, Valley Vet Supply; 37, Travel Ink/Getty Images; 41, Elemental-
Imaging/iStockphoto; 44 top, Edgar Fahs Smith Memorial Collection;

44 bottom, Fotolia; and 48, Library of Congress.

Chapter 3 Page 56, Thomas Deerinck, NCMIR/Photo Researchers, Inc.; 57,
Argonne National Laboratory; 58, Pascal Goetgheluck/Photo Researchers, Inc.;
60, iStockphoto; 69, iStockphoto; 70, MARKA/Alamy; 71, Bio-Rad
Laboratories, Inc.; 73 top, REUTERS/William Philpott WP/HB; 73 bottom,
AFP Photo/Newscom; and 78, Bettmann/CORBIS.

Chapter 4 Page 85, Shutterstock; 86, Swiss Institute of Bioinformatics; 88,

Lisa A. Shoemaker; 89 top, Bror Strandberg; 89 bottom, Hulton Archive/Getty
Images; 93, Custom Life Science Images/Alamy; 94, Bettmann/ Corbis; 95,
Julian Voss-Andreae; 108, From Kiihner et al., “Proteome Organization in a
Genome-Reduced Bacterium” Science 27 Nov 2009 Vol. 326 no. 5957
pp. 1235-1240. American Association for the Advancement of Science.; 109,
Howard Ochman; 111, From Butland et al., “Interaction network containing
conserved and essential protein complexes in Escherichia coli,” Nature 433
(2005), 531-537; 113, National Library of Medicine; 117, Laurence A. Moran;
119, Easawara Subramanian, http://www.nature.com/nsmb/journal/v8/n6/full/
nsb0601_489.html; 121, Danielle Anthony; 122, SSPL/The Image Works; 123,
Janice Carr/Centers for Disease Control; 126, Ed Uthman, licensed via Creative
Commons http:// creativecommons.org/licenses/by/2.0/; and 127, Julian
Voss-Andreae.

Chapter 5 Page 135, Dorling Kindersley; 136, Jonathan Elegheert; 137, Michael
P. Walsh/IUBMB; 138, Leonardo DaVinci; 142 top, Rockefeller Archives
Center; 142 bottom left, University of Pittsburgh, Archives Service Center;

142 bottom right, Laurence A. Moran; and 149, AP Photo/Paul Sakuma.

Chapter 6 Page 167, Ronsdale Press, photo copyright Dina Goldstein; 174,
Bettmann/CORBIS; 183, Paramount/Photofest; and 186, Shutterstock.

Chapter 7 Page 198, Shutterstock; 200, Library of Congress; 204, Heath
Folp/Industry & Investment NSW; 209, History Press; 212, Christian Heintzen,
University of Manchester; 214, iStockphoto; 215, John Olive; 216, Stephanie
Schuller/Photo Researchers, Inc.; 219 left, Meg and Raul via Flickr/CC-BY-2.0
http://creativecommons.Org/licenses/by/2.0/deed.en 219 right, and 220,
Shutterstock; and 223, both, ©® The Nobel Foundation.

Chapter 8 Pages 227, 239, 240, Shutterstock; 244 top, Image Source/ Alamy;

244 bottom, Jack Griffith; 245, Jakob Jeske/Fotolia; 246, Jens Stougaard; 247
top, Eric Erbe, Christopher Pooley, Beltsville Agricultural Resear ch/USDA;

247 bottom, Robert Hubert, Microbiology Program, Iowa State University; and
252, Christine Ortlepp.

Chapter 9 Page 258, imagebroker/ Alamy; 262 top, Steve Gschmeissner/Photo
Researchers, Inc.; 262 bottom, Shutterstock; 268 bottom, Shutterstock; 270,
John Ross; 273 top, Professors Pietro M. Motta 8c Tomonori Naguro/Photo Re-
searchers, Inc.; 273 bottom, Biophoto Associates/Photo Researchers, Inc.; 277,
Lisa A. Shoemaker; 278 bottom, Julie Marie/Fotolia; 284 top, M.M. Perry; and
284 bottom, Shutterstock.

Chapter 10 Page 294, Quade Paul, Echo Medical Media; 296, Charles Boone,
From Costanzo et al. “The Genetic Landscape of a Cell” Science 327 \
(2010):425-432; 297, Roche Applied Science; 303, Shutterstock; 305 top,
University of Edinburgh/Wellcome Images; 305 bottom, Biophoto
Associates/Photo Researchers, Inc.; and 312, National Library of Medicine.

Chapter 11 Page 325, Barton W. Spear — Pearson Education; 331 left, Super-
Stock, Inc;. 331 right, Bettmann/CORBIS; 336, Warner Bros./Photofest; 341,
ChinaFotoPress/Zuma/ICON/Newscom; and 349, dreambigphotos/Fotolia.

Chapter 12 Page 359, CBS/Landov; 369, United States Postal Service; 370 top,

A. Jones/Photo Researchers, Inc.; 370 bottom, Laura Van Niftrik; and 375,

Tim Crosby/Getty Images.

Chapter 13 Page 386, Science Photo Library/Photo Researchers, Inc.; 387,

From Zhou, Z.H. et al. (2001) Proc. Natl. Acad. Sci. USA 98, pp. 14802-14807;
390 top, From Zhou, Z.H. et al. (2001) Proc. Natl. Acad. Sci. USA 98, pp.
14802-14807; 390 bottom, NASA; and 396, 401, Shutterstock.

Chapter 14 Page 417 top and left, Shutterstock; 417 bottom, Dirk Freder/
iStockphoto; 419 top, Lisa A. Shoemaker; 419 middle and bottom, Shutterstock;
420 top Roberto Danovaro; 420 left, Milton Saier; 426, Michael Radermacher;
433, Alexander Tzagoloff; and 438, NASA/Sandra Joseph and Kevin O’Connell.

Chapter 15 Page 443, Mary Ginsburg; 444, Arizona State University — Plant
Bio Department; 447 top, Makoto Kusaba; 447 bottom, Shutterstock; 448 top,
CHINE NOUVELLE/SIPA/Newscom; 448 bottom, Robert Lucking; 452, Niels
Ulrik Frigaard; 457, Michelle Liberton, Howard Berg, and Himadri Pakrasi, of
the Donald Danforth Plant Science Center and of Washington University,

St. Louis; 458 top, Andrew Syred/Photo Researchers, Inc.; 458 bottom, NSF
Polar Programs/NOAA; 459, Lisa A. Shoemaker; 462, Lawrence Berkeley
National Laboratory; 468, Shutterstock; 469 top, From Bhattacharyya et al,
“The wrinkled-seed ...” Cell, Vol 60, No 1, 1990, pp 115-122; 469 middle, Peter
Arnold/Photolibrary; 469 bottom, Fotolia; 470, From David F. Savage et al.,
“Spatially Ordered Dynamics of the Bacterial Carbon Fixation Machinery,”
2011. American Association for the Advancement of Science; 471 top,

AP Photo/Charlie Neibergall; and 471 bottom, Shutterstock.

Chapter 16 Page 475, Kennan Ward/Corbis; 486, Shutterstock; 490 top,
Bettmann/CORBIS; 490 bottom, Hulton Archive/Getty Images; 493, Environ-
mental Justice Foundation, Ltd.; 495 top, David Leys, Toodgood et al., 2004;

495 bottom, Eric Clark/Molecular Expressions; 501 top, Shutterstock; 501
bottom, Steve Gschmeissner/SPL/ Alamy; 504, Donald Nicholson/IUBMB; 506,
Shutterstock; and 507, Robin Fraser.

Chapter 17 Page 515 top, NASA Visible Earth; 515 bottom, NOAA; 516, Inga
Spence/Photo Researchers, Inc.; 531, Shutterstock; 532, iStockphoto.com; 534,
National Library of Medicine; and 540, U.S Air Force photo/Staff Sgt Eric T.
Sheler.

Chapter 18 Page 552, G. Robert Greenberg; 554, National Library of Medicine;
561, Peter Reichard; 564, Shutterstock; and 568, Fotolia.

Chapter 19 Page 574, National Cancer Institute; 581, SSPL/The Image Works;
587, Andrew Paterson/ Alamy; 589, Lisa A. Shoemaker; 591 both, Ulrich K.
Laemmli; 597 top left, 597 top right, Lisa A. Shoemaker; 597 middle, Stanford
University School of Medicine; and 597 bottom, Steve Northup/Time&Life
Images/ Getty Images.

Chapter 20 Page 603 top, John Cairns; 603 bottom left, David S. Hogness; 603
bottom right, Regional Oral History Office, The Bancroft Library, University of
California, Berkeley; 613 both, Timothy Lohman; 615, From Structure, 6,

Dec. 2008 Copyright Elsevier. Original artwork by Glass Egg Design, Jessica
Eichman, www.glasseggdesign.com; 618, Lisa A. Shoemaker; 619, David
Bentley; 627 top, Laguna Design/Photo Researchers, Inc.; 627 bottom, Paul
Sabatier/ Art Life Images/Superstock; 628 top, James Kezer/Stanley Sessions;

628 bottom, Dr. L. Caro/Photo Researchers, Inc; 630, Institute of Molecular-
biology and Biophysics, From Yamada et al., Molecular Cell Vol 10 p 671
(2002). Figure 4b (right), with permission from Elsevier.; and 630, Vanderbilt
University, Genes and Development. From Wang et al. BASC, a super complex
of BRCA1 -associated proteins involved in the recognition and repair of
aberrant DNA structures. Vol. 14, No. 8, pp. 927-939, April 15, 2000 Fig 3M.

Chapter 21 Page 634, Marc Gantier/Getty Images; 636, From Murakami.et al.,
Science 296: 1285-1290 (2002) Fig5A (left) American Association for the
Advancement of Science; 638, Oscar L. Miller, Jr.; and 651, Lisa A. Shoemaker.

Chapter 22 Page 666, National Security Agency; 667, US Navy Office of
Information; 675, David Goodsell; 677, Stanford University School of
Medicine; 681, Oscar L. Miller, Jr.; and 692, H. H. Mollenhauer/USDA.

767

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Index

In this index, the page numbers listed indicate
tables (with a T added to that page number) and
figures (with an F added to that page number).

A-DNA, 585-586F
ABO blood group, 250-25 IF
absorption spectrum of DNA, 584-585F
acceptor stem, 668F
accessory pigments, 447-448F
acetaldehyde, lyases catalyzation, 137
acetaminophen, structure of, 486F
acetate, gluconeogenesis precursor, 362-363
acetic acid (CH 3 COOH), 45
buffer range of, 50F
dissociation of, 45
pH and, 45, 47, 50F
titration of, 47F
acetyl CoA, 315-316, 387-394
cholesterol and, 488
citric acid cycle reactions, 385, 387-394
isopentenyl diphosphate conversion from, 488
nucleotidyl group transfer, 315
oxidation of, 385, 391-394F
pyruvate, conversion from, 385, 387-391
thioester hydrolysis, 316
acetylcholinesterase, 134F
acid-base catalysis, 168-169
acid solutions, 42-49F

base solutions combined with, 47-48
base solutions dissociated from, 44-45
dissociation constant, K a , 44-48T
Henderson-Hasselbach equation for, 46-47
ionization and, 42
pH scale for, 43F, 49
parameter value, p K a , 45-48T
titration, curves for, 47-48F
weak, 44-49

aconitase, citrus cycle reactions, 3 96-3 9 7F
actin filaments, 23F
activation energy, G*, 14F, 165F
activator ions, 196

active membrane transport, 280-283F
acute lymphoblastic leukemia treatment, 52 1
acyl, general formula of, 5F
acyl carrier protein (ACP), 1 1 IF, 204-206F
acyl CoA transport into mitochondria, 497-498
adenine (A), 8-9F, 310-31 IT, 551F
adenosine deaminase, 181-182F
adenosine 5-monophosphate (AMP), 550-55 IF
adenosine triphosphate (ATP), 8-9F, 198-199F,
308-315,417-442

active membrane transport, 282-283F, 435-436
/7-oxidation, generation from, 498-499
citric acid cycle reactions, 405-406F
coenzyme metabolic property, 198-199F
cyclic adenosine monophosphate (cAMP),
287-288F

electron transport and, 417-442
eukaryotic mitochondria and, 2 1
Gibbs free energy change, AG, 308-312
hexokinase reactions, 326-327, 328F, 330F
high energy bond, ~, 3 1 1
hydrolysis, 308-312

electrostatic repulsion, 309
metabolically irreversible changes, 308-312

resonance stabilization, 310
solvation effects, 309-310
metabolic changes, 198-199F, 304, 308-315
nucleotide metabolic reactions, 55 IF
nucleotidyl group transfer, 315F
phosphofruktokinase- 1 (PFK-1) regulation by,
345-346F

phosphoryl group transfer, 312-315

photosynthesis photosystems and, 459-460F

production of, 314-315F

reduced coenzyme production of, 405-406F

structure of, 8-9F

synthase, 433-435F, 456, 459-460F

synthesis of, 417-442

ATP synthase catalysis, 433-435F
chemiosmotic theory, 420-423
mitochondria, 418-420F
mitochondrial membrane transport, 435-436
NADH shuttle mechanisms in eukaryotes,
436-439F

P/O (phosphorylated/ oxygen) ratio, 436
proton leaks and heat production, 435
protonmotive force, 421-420F
superoxide anions, 440-441
adenylyl cyclase signaling pathway, 287-288F
adenylyl kinase (pig), 105F
affinity chromatography, 70
aggrecan, 245-246

aggregation from protein folding, 119
Agre, Peter, 280
Agrobacterium sp., 528
alanine (A, Ala), 56, 59F, 64T
catabolism of, 535
gluconeogenic precursor, 361
glucose- alanine cycle, 36 IF
ionization of, 64-65F
isomerases catalyzation, 137-138
nomenclature, 56, 64T
pyruvate, conversion from, 36 IF
structure and properties of, 56, 59F
synthesis of, 521-523F
titration of, 64-65F
transferases catalyzation, 136-137
alcohol groups with side chains, 60-61
alcohols, 5F

cyclization of monosaccharides and reactions of,
230-23 IF

general formula of, 5F
solubility in water, 35T
aldehyde, general formula of, 5F
aldohexoses, 229F
aldolase cleavage, 330-332F
aldopentoses, 229F
aldoses, 228-234F

cyclization of, 230-234F
epimers, 230

Fischer projections of, 228-230F
structure of, 228-230F
aldotetroses, 229F
aliphatic R groups, 59
alkaline hydrolysis, 591-592F
alkaptonuria, 544
allose, 229F

allosteric enzymes, 153-158F

concerted (symmetry) model for, 156-157F
phosphofructokinase, 154-155F

properties of, 155-156F
regulation of enzyme activity using, 153-158
sequential model for, 157-158F
allosteric protein interactions, 127-129F
allosteric regulation of eukaryotic ribonucleotide
reductase, 56 IT
allysine residues, 12 IF
or- carbon atom, 56
or-globin subunits, 122-123F
or helix proteins, 94-97F, 98-99
amphipathic, 95-97A
P strand and sheet connections, 98-99F
collagen type III triple helix, 1 19F
left-handed, 119-120F
leucine zipper, 96-97 A
membranes, 270-271F
protein conformation of, 94-9 7F
right-handed, 94-95F
rotation of, 95
side chains in, 95
3 10 helix compared to, 96-97F
a- ketoglutarate, transferases catalyzation, 56
or-ketoglutarate dehydrogenase complex, citrus
cycle reactions, 398-399F
or subunits, RNA transcription, 641-642T
or- tocopherol (vitamin E), 218F
or//?barrel, domain fold, 106F
a ifh tetramer (insulin), 290-29 IF
altrose, 229F
amide linkages, 4-5F
amino acid metabolism, 514-549
ammonia assimilation, 518-519
glutamate and glutamine
incorporation, 518F
transanimation reactions, 518-519F
catabolism, 534-542

alanine, asparagine, aspartate, glutamate,
and glutamine, 535

argenine, histidine, and proline, 535-536F
branched chain amino acids and, 537-539F
cysteine, 540-54 IF
glycine and serine, 536-537F
lysine, 542F

methionine conversion and, 539-540F
threonine, 537-538F
tyrosine, 541-542F
diseases of, 544
essential amino acids, 529T
functions of, 514-515
nitrogen cycle, 515-517F
nitrogen fixation, 515
nitrogenases, 516-517
nonessential amino acids, 514, 529T
precursors, 529-532

glutamate, glutamine, and aspartate, 529
lignin from phenylalanine, 531-532F
melanin from tyrosine, 531, 533F
nitric oxide from arginine, 530-53 IF
serine and glycine, 529-530F
protein turnover, 531-533
renal glutamine metabolism, 547-548
synthesis of amino acids, 520-529

alanine, valine, leucine, and isoleucine,
521-523F

aspartate and asparagine, 520-52 IF
citric acid cycle, 520F

769

770 INDEX

amino acid metabolism ( Continued )

glutamate, glutamine, arginine, and
proline, 523F
histidine, 527F

lysine, methionmine, and threonine, 520-522F
phenylalanine, tyrosine, and tryptophan,
524-527F

serine, glycine, and cysteine, 523-525F
urea cycle, conversion of ammonia to urea,
542-547

amino acids, 6F, 55-84
a-c arbon atom, 56
active sites of enzymes, 168T
catabolism of, 519, 534-542
catalytic functions of residues, 166-168T
chromatographic procedure for, 73-74F
common types of, 58-62

alcohol groups with side chains, 60-61
aliphatic R groups, 59
aromatic R groups, 59-60
derivatives, 62-63
hydrophobicity of side chains, 62
negatively charged R groups, 62
positively charged R groups, 61-62
sulfur- containing R groups, 60
defined, 56

evolution and ancestors from, 57-58, 79-8 IF

free-energy change of transfer for, 63T

glucose precursors, 360-361

hydrolysis for analysis of, 73-74F

hydropathy scale, 62T

ionization of, 63-67

molecular weight of, 74-75T

nomenclature, 56-58, 6 IF, 64T

peptide bonds, 67-68

pK a values, 168T

protein composition with, 67-68, 73-74T
protein purification and analysis, 68-73
racemization, 58
residues, 67-68F, 74-75F
RS system configuration, 6 IF
sequencing, 68, 74-8 IF
side chains, 56, 59-62
site-directed mutagenesis, 167
structure of, 6F, 56-62F
abbreviations for, 58-59F
ball- and- stick model of, 56-5 7F
mirror-image pairs, 57F
numbering conventions, 56F
titration of, 64-65F
amino sugars, 235-236, 237F
aminoacyl-tRNA, 670-673

binding sites, 671-672F, 675F, 677F
docked at A site, 675, 677F, 680-682F
elongation factors and docking of, 680-682F
ribosome binding sites, 675, 677F
synthetases, 670-673F

proofreading for errors in, 673
protein synthesis and, 670-673F
reaction of, 670-672F
specificity of, 671-673F
substrate-binding sites, 677F
aminoimidazole carboxamide ribonucleotide
(AICAR), 553F

aminoimidazole ribonucleotide (AIR), 553F
aminoimidazole succinylocarboxamide
ribonucleotide (SAICAR), 553F
ammonia (NH 3 ), 45, 518-519
assimilation, 518-519
conversion to urea, 542-547
dissociation for formation of, 45
enzyme transfer from glutamate, 558
glutamate and glutamine incorporation, 518F
transanimation reactions, 518-519F
urea cycle, 542-547F
ammonium ions, general formula of, 5F
amphibolic pathways, 407-409

amphibolic reactions, 295
amphipathic helix, 95-97A
amphipathic molecules, 36
amplification, 285
DNA, 615-616
signal pathways, 285
amylase, 242F
amylopectin, 241-242F
amyloplasts, 469
amylose, 24 IF
Anabaena spherica, 305F
anabolic (biosynthetic) reactions, 294-295F,
302-303F

anaerobic conversion, 339-340F
Anfinsen, Christian B., 112-113
angstrom (A), units of, 26
anionic forms of fatty acids, 258T
anomeric carbon, 231
anomers, 23 1

antenna chlorophylls, 446-447F
anti conformation of nucleotides, 577-578F
antibiotic inhibition of protein synthesis, 686
antibody binding to specific antigens, 129-130F
anticodon arm, 668-669F
anticodons, 668-67 IT
base pairing, 669-670T
defined, 668

wobble position of, 670-67 IF
antigens, antibody binding to, 129- DOF
antiparallel /? sheets, 97-98F
antiparallel DNA strands, 581-583
antiport, membrane transport, 28 IF
apoptosis, 534
aquaporin, 280F
Arabidopsis thalianna , 93
arabinose, 229F

L-arabinose-binding protein, 105F
arginine (R, Arg), 61-62F
catabolism of, 535-536F
nitric oxide synthesis from, 530-53 IF
nomenclature of, 64T
structure of, 61-62F
synthesis of, 523F
urea cycle and, 543F, 545-546F
arginine kinase, 190-192F
aromatic R groups, 59-60
arsenate (arsenic) poisoning, 336
arsenite (arsenic) poisoning, 336
ascorbic acid (vitamin C), 209-211
asparagine (N, Asn), 62F

acute lymphoblastic leukemia treatment, 521
catabolism of, 535
nomenclature, 64T
structure of, 62F
synthesis of, 520-52 IF
aspartame, 68F, 240
aspartate (D, Asp), 62F
catabolism of, 535
gluconeogenic precursor, 361
malate-aspartate shuttle, 348F
metabolic precursor use, 529
nomenclature, 64T
structure of, 62F
synthesis of, 520-52 IF
urea cycle and, 543F, 545-546F
aspirin, structure of, 486F
association constant, K a , 109-1 10F
atmospheric pollution, photosynthesis
and, 457

ATP, see adenosine triphosphate (ATP)

ATP synthase, 43 3-43 5F

binding change mechanism, 434-43 5F

chloroplasts, 459-460F

cytochrome complexes, 456

electron transfer from, 456

electron transport, complex V, 433-435F

photosynthesis and, 456, 459-460F

rotation of molecules, 434-435
structure of, 433F
attenuation, 688-689F

audioradiograph of replicating chromosome, 603F
autophosphorylation, 290
autotrophs, 302-303
Avery, Oswald, 3, 573

Azotobacter vinelandii nitrogenase , 516-517F

B-DNA, 582-584F

bacteria, 246-248. See also Escherichia coli ( E . coli )
citric acid cycle and, 411-414
Entner-Doudoroff (ED) pathway, 351-352F
forked pathway, 412-413F
gloxylate pathway, 41 1-41 2F
Gram stain for, 247F
intestinal, 216F

metabolism and adaptation of, 295-296
penicillin, 247-248F
peptidoglycans, 246-248F
polysaccharide capsules, 247
Staphylococcus aureus (S. Aureus ), 76, 247-248F
bacterial DNA, 3, 590
bacterial enzymes, 364F
bacterial flagellum, 109F
bacterial photosystems, 448-458
coupled, 453-455T
cytochrome bf complex, 453-455F
electron transfer in, 449-453
Gibbs free energy change, AG, 455-457
green filamentous bacteria, 448, 452F
internal membranes, 457
photosystem I (PSI), 448, 450-453F
photosystem II (PSII), 448-450F
purple bacteria, 448-450F
reaction equations, 450T, 452T, 455T
reduction potentials, 455-457F
bacterial reaction center (BRC), see photosystems
bacterial transducers, 285-286
bacteriophage MS2 capsid protein, 107F
bacteriorhodopsin, 270-271F, 461
ball-and-stick models, 56-57F
amino acids, 56-57F
DNA, 582-584F

monosaccharide (chiral) compounds, 228F, 235F
Barnum, P. T„ 200
Bascillus stearothermophylus , 402
Bascillus subtilis, 186
base composition of DNA, 579T
base pairing, 604-606, 669-671
DNA, 604-606
protein synthesis, 668-67 IF
Watson-Crick, 668-670F
wobble positions of anticodon and codon,
670T-671F

base solutions, 42-43F, 47-48F
acid titration using, 47-48F
dissociated from acid solutions, 44-45
Henderson-Hasselbach equation for, 47-48
ionization and, 42
pH scale for, 43F
Beadle, George, 212, 634
/^barrel, domain fold, 106F
/^barrel protein membranes, 271-272F
p- carotene, 217F, 447F
/2-globin subunits, 122-123F
/?helix, domain fold, 106F
/2-meander motif (structure), 100-101F
p- oxidation, 494-501

acyl CoA transport into mitochondria, 497-498

ATP generation from, 498-499

fatty acids, 494-501

lipid metabolism and, 494-501

odd- chain fatty acids, 499-500

trifunctional enzymes and, 498

unsaturated fatty acids, 500-501

Index 771

/2-sandwich motif (structure), 100-10 IF
P strands and sheets, 97-99F
or helix connections, 98-99F
antiparallel sheets, 97-98F
P turns, 99F

hydrophobic interactions, 98
loops, 98

parallel sheets, 97-98F

pleated sheet, 97-98

protein conformation of, 97-99F

residues and, 99F

reverse turns, 99

turns, 99F

pap unit motif (structure), 100F
bicarbonate production by renal glutamine
metabolism, 547-548
bidirectional DNA replication, 602-603F
bile salts, 505F

binding. See also oxygen binding; substrates
aminoacyl-tRNA sites, 671-672F,

675F, 677F

cap binding protein (CBP), 679

change mechanism, ATP synthase, 434-43 5F

DNA fragments, 609-61 IF

hormones, 286-288

protein synthesis, 671-672F, 675F,

677-679F
biochemistry, 1-27
biopolymers, 4-10
cells, 17-26

E. coli , 17F, 23-24, 26F
eukaryotic, 18-23F
living, 23-26
prokaryotic, 17-18F
chemical elements of life, 3-4
defined,

energy, life and, 10-15
evolution and, 15-17
macromolecules, 4-10
lipids, 9

membranes, 9-10
nucleic acids, 7-9F
polysaccharides, 6-7F
proteins, 6

multidisciplinary nature of, 26
special terminology of, 26-27
20th century science and, 2-3
units for, 26-27T

bioenergetics, 11 .See also ATP; metabolism;
thermodynamics

biological functions, 55-56, 119-129
amino acid metabolism diseases, 544
antibody binding to specific antigens,

129-130

blood plasma, 33F, 35F, 51-52F

cancer DCA inhibitors, 408F

cartilage structure, 245-246F

coronary heart disease and lipoprotein lipase, 507

diabetes mellitus (DM), 381, 51 1

dietary requirements and fatty acids, 261

genetic defects, 265-266

gout, 569

hyperactivity, 359

intestinal bacteria, 216F

lactate buildup, 341

lactose intolerance, 350

Lesch-Nyhan syndrome, 569

gout, 569

lysosomal storage diseases, 492F
liver metabolic functions, 344-345F,

379-380F

mucin secretions, 252F

oxygen binding to myoglobin and hemoglobin,
123-129

proteins and, 55-56, 119-129
scurvy, ascorbic acid and, 209-210
sweetness receptors, 240

vitamin deficiency, 198 T, 209-210, 214, 215
biological membranes, 9, 269-275.

See also membranes
biopolymers, see polymers

biosynthetic (anabolic) pathways, 302-303
biotin, 21 1-2 12F

2,3-bisphospho-D-glycerate (2,3BPG),
127-128F

1.3 Hsphosphoglycerate, 334F

2.3 Hsphosphoglycerate, 335-337F
bisubstrate enzyme reactions, 147-148F
blood, 33F,35F,250-251F

ABO group, 250-25 IF

2,3 Hsphosphoglycerate in, 335F
buffer capacity, 51-52F
glycolysis reactions, 335
plasma, 33F, 35F, 51-52F
properties of, 33F, 35F
boat conformations, 235F
Bohr effect, 128F
Boyer, Herbert, 597
Boyer, Paul D., 223, 434
branched chain amino acids, 537-539F
breast cancer and DNA repair, 630
Briggs, George E., 141
Buchanan, John (Jack) M., 551, 554
Buchner, Eduard, 2, 331
buffered solutions, 50-52F
acetic acid, 50F
blood plasma, 51-52F
capacity and pK a , 50-52FT
carbonic acid, 5 IF
pH and, 50-52F
preparation of, 50

C-terminus (carboxyl terminus), 68, 76F

C 3 pathway, see Calvin cycle

C 4 pathway, 469-47 IF

Caenorhabditis elegans , 296

Cahill, George, 380

calcium (Ca), 3

calories (cal), units of, 26

calorimeter, 13F

Calvin, Melvin, 462

Calvin cycle, 443, 461-467F

carbon dioxide (C0 2 ) fixation, 461-467,
469-472

NADPH reduction, 466-467
ribulose 1,5-frzsphosphate, 465-466F
rubisco (rubilose 1,5-frisphosphate

carboxylase- oxygenase), 462, 464-466F
stages of, 462F

oxygenation, 465-466F
reduction, 466-467
regeneration, 466-467F
cancer drug inhibition, synthesis for, 564
cap binding protein (CBP), 679
cap formation, mNRA, 658-659F
capsaicin, 284F
capsule, polysaccharide, 247
carbamate adducts, 129F
carbamoyl phosphate, urea cycle and, 543F,
545-546F

carbamoyl phosphate synthetase, 558F
carbocation, 164
carbohydrates, 227-255
defined, 227
disaccharides, 236-239

glycosidic bonds in, 236-238F
structures of, 237-239F
sugars, 238-239
glucosides and, 236-239, 24 IF
nucleosides and, 239, 241F
glucosides, 236-239, 24 IF
glycoconjugates, 244-252
glycoproteins, 248-252F

peptidoglycans, 246-248F
proteoglycans, 244-246F
monosaccharides, 227-236
aldoses, 228-234F
ball- and- stick models of, 228F
chiral compounds, 228-230F
conformations of, 234-235F
cyclization of, 230-234
derivatives of, 235-236F
epimers, 230

Fischer projections of, 228-232F
Haworth projections of, 232-235F
ketoses, 228-234F
trioses, 226

oligosaccharides, 227, 248-252F
polysaccharides, 227, 240-244
cellulose, 243F
chitin, 244F
glycogen, 240-243F
heteroglycans, 240
homoglycans, 240
starch, 240-242F
structure of, 240-24 IT
carbolic acid, general formula of, 5F
carbon (C), 3

glycolysis reactions, 333-334F
carbon dioxide (C0 2 ), lyases catalyzation, 137
carbon dioxide (C0 2 ) fixation, 461-467
bacteria compartmentalization, 469
C 4 pathway, 469-47 IF
Calvin cycle, 443, 461-467F
carboxysomes, 469-470F
crassulacean acid metabolism (CAM),
471-472F

NADPH reduction, 466-467
ribulose 1,5-frzsphosphate, 465-466F
rubisco (rubilose 1,5-frisphosphate

carboxylase- oxygenase), 462, 464-466F
carbonic acid, buffer capacity of, 51F
carbonic anhydrase, 197F
carbonyl, general formula of, 5F
carboxyaminoimidazole ribonucleotide
(CAIR), 553F

carboxylate, general formula of, 5F
carboxysomes, 469-470F
carotenoids, 447-448F
cartilage structure, 245-246F
cascade amplification of signal pathways, 285
catabolic reactions, 295F, 303-304F. See also
glycolysis
glucose, 325-354
metabolic pathways, 303-304F
NADH, 304
catabolism, 534-542

alanine, asparagine, aspartate, glutamate, and
glutamine, 535

amino acid metabolism and, 534-542
argenine, histidine, and proline, 535-536F
branched chain amino acids and, 537-539F
cysteine, 540-54 IF
glycine and serine, 536-53 7F
lysine, 542F

methionine conversion and, 539-540F
purine, 565-568
pyrimidine, 568-570
threonine, 537-538F
tyrosine, 541-542F
catalysis, 166-171, 175-182
acid-base, 168-169
amino acid residues and, 166-168T
catalytic residue frequency distribution, 168T
chemical modes of, 166-171
covalent, 169-170F

diffusion- controlled reactions, 171-175
enzymatic modes, 175-182
induced fit, 179-180
proximity effect, 176-178F

772 INDEX

catalysis ( Continued )

transition-state stabilization, 176, 180-182F
weak binding and, 176, 179-179F
enzyme mechanism of, 166-171, 175-182
ionizable amino acid residue functions,
166-168T

pH effects on enzymatic rates, 170-172F
p K a values of ionizable amino acids, 168T
RNA polymerase, 637-638F
serine proteases and modes of, 185-188
substrate binding and, 171-172T, 175-182F
catalysts, 2, 113-114, 134, 136-138
defined, 134

denaturation reduction from, 113-114
hydrolase enzymes, 137
isomerases enzymes, 137-138
ligases enzymes, 138
lyases enzymes, 137
oxidoreductase enzymes, 136
protein structures, 113-114
regulation of enzyme activity, 153-158
transferases enzymes, 136-137
catalytic activity, 89
catalytic constant, k cat , 143-145
catalytic proficiency, 144-147T
catalytic triad, 185F
cellobiose, 237-238, 239F
cells, 17-26

cytosols, 23, 26F
E. coli, 17F, 23-24, 26F
diffusion in, 34F
eukaryotic, 18-23F
living, 23-26
prokaryotic, 17-18F
structure of, 17-23
solubility and concentrations of, 34F
cellular pathways, 302-304
cellulose, 243F
cellulose, 7-8F

Celsius scale (°C), units of, 26-27
Central Dogma, 3
cerebrosides, 265, 266F
ceremide, 264, 265F
chain elongation, 603, 679-684

DNA polymerase replication, 604-606F
protein synthesis translation, 673-674, 679-684
aminoacyl-tRNA docking sites for, 680-68 IF
elongation factors, 680-68 IF
microcycle steps for, 679-684F
peptidyl transferase catalysis, 681-682F
ribosomes and, 673-674
translocation of ribosome, 682-684F
RNA polymerase catalyzation, 63 6-63 7F
chair conformations, 189-190F, 235F
Chance, Britton, 420
Changeaux, Jean-Pierre, 157
channels for (animal) membrane transport,
279-280F
chaotropes, 36

chaotropic agents for denaturation, 111
chaperones, see molecular chaperones
Chargaff, Erwin, 579
charge-charge interactions, 37, 117, 584
chemiosmotic theory, 420-423
chemoautotrophs, 303, 439-440
chemoheterotrophs, 303
chemotaxis, 284
chiral atoms, 56-57
chiral compounds, 228-230F
chitin, 244F

Chlamydomonas sp., 458
chloride (Cl), 3
chlorophylls, 444-447F
antenna, 446-447F
photon (energy) absorption, 445-446
resonance energy transfer, 446

special pair, 446-447F
structure of, 444-445
chloroplasts, 21-22F, 458-460F
ATP synthase, 459-460F
cyanobacteria evolution of, 459
eukaryotic cell structure and, 20F, 21-22F
organization of, 459-460F
photosynthesis and, 22
structure of, 458-459F
cholecalciferol (vitamin D), 218-219F
cholesterol, 266-268

isoprenoid metabolism and, 490, 493-494F
level regulation, 493
lipid bilayers, 277-278F
lipid metabolism and, 488, 490-494
membrane fluidity and, 277-278F
steroids and, 266-268
synthesis of, 488, 490-494
chromasomal DNA replication, 602-603
chromatin, 588-591

bacterial DNA packaging, 590
higher levels of, 590
histones, 588-590F
nucleosomes and, 588-591
packing ratio, 588

RNA eukaryotic transcriptions and, 649
chromatography, 69-70F, 73-74F
amino acid analysis, 73-47F
techniques, 69-70F
chymotrypsin, 76-77F, 183-188F
Ciechanover, Aaron, 533
cis conformation, 9 IF, 93, 258, 259F
cis/trans isomerization, 93, 104F
cistine, formulation of, 60F
citrate synthase, citrus cycle reactions, 385F,
394-396F

citric acid cycle, 303-304, 326F, 385-416
amphibolic pathways, 407-409
ATP production, 405-406F
bacteria and, 411-414
coenzyme reduction, 405-406F
energy production in, 405T
enzymatic reactions of, 392
enzyme reactions, 386, 394-402
aconitase, 3 96-3 9 7F

or-ketoglutarate dehydrogenase complex,
398-399F

citrate synthase, 394-396F
conversion of from another, 402F
fumarase, 401

isocitrate dehydrogenase, 397-398F
malate dedrogenase, 401-402
succinate dehydrogenase complex, 399-40 IF
succinyl synthetase, 398-400F
eukaryotic cells and, 385
evolution of, 412-414
forked pathways, 413F
gloxylate pathway, 409-412
glucose synthesis from, 326F
glycolytic pathway, 408
history of, 385-386
metabolic pathway, 303-304
oxidation of acetyl CoA, 385, 391-394
prochiral substrate binding, 397
pyruvate conversion to acetyl CoA, 385, 387-391
pyruvate entry into mitochondria, 402-405F
regulation of, 406-407
cleavage, 76-77F, 112F, 163-164
bonds, 112F, 163-164
carbocation, 164
enzyme reactions and, 163-164
free radicals, 164
hydrolysis, 592F, 594F
nuclease sites, 592F

proteins by cyanogen bromide (CNBr), 76-77 F
RNA, 594F

Cleland, W. W., 147
cobalamin (vitamin B 12 ), 215-216F
codons, 665-670T
anticodons, 668-67 IF
base pairing, 669-670T
defined, 665
genetic code, 665-668F
initiation, 667, 675-679F
mRNA reading frames, 666-667F
protein synthesis and, 665-684
RNA translation and, 675-679F
synonymous, 667
termination (stop), 667, 682, 684
translation of in chain elongation, 679-684F
wobble positions, 670-67 IF
coenzymes, 196-226, 316-321

acyl carrier protein (ACP), 204-206F
adenosine triphosphate (ATP), 198-199F,
405-406F

ascorbic acid (vitamin C), 209-211
biotin (vitamin B 7 ), 21 1-21 2F
citric acid cycle, 405-406F
cobalamin (vitamin B 12 ), 215-216F
coenzyme A, 204-206F
cofactors, 196F
cosubstrates, 197-199
cytochromes, 221-222F
electron transfer for free energy, 319-320
energy conservation from, 316-320
flavin adenine dinucleotide (FAD), 204-205F
flavin mononucleotide (FMN), 204-205F
Gibbs free energy change, AG, 317-319
half-reactions, 317-319T
inorganic cations, 197
lipid vitamins, 2 1 7-2 1 9F
lipoamide, 216-217F
mechanistic roles, 199T
metabolic roles of, 198-200T
metal- activated enzymes, 197
metalloenzymes, 197
NADH reactions, 319-320
nicotinamide adenine dinucleotide (NAD),
196F, 200-203F

nicotinamide adenine dinucleotide phosphate
(NADP), 200-202F
nobel prizes for, 223
nucleotides, 198-199
oxidation-reduction, 221F, 316-320
prosthetic groups, 197, 205-206F
proteins as, 22 1

pyridoxal phosphate (PDP), 207-209F
reactive center, 196
reduced, 316-320, 405-406F
reduction potential, 317-319T
riboflavin, 204-205F
tetrahydro folate, 213-214F
thiamine diphosphate (TDP), 206-207F
ubiquinone (coenzyme Q), 2 19-22 IF
vitamins, 196, 198-199T
co factors, 196F, 425
Cohen, Stanley N., 597
coiled-coil motif (structure), 100F
collagen, 11 9-12 IF

covalent (bond) cross links in, 120- 12 IF
interchain hydrogen bonding in, 120F
protein structure, study of, 11 9- 12 IF
residue formation and, 120-12 IF
Schiff bases, 12 IF
type III triple helix, 1 19F
column chromatography, 69-70F
compartmentation, 304-305
complementary base pairing, double-helix DNA,
582-583F

concanavalin A (Jack bean), 104F
concerted (symmetry) model for enzyme
regulation, 156-157F

Index 773

configurations versus conformations, 234
conformational changes from oxygen binding,
124-126F

conformations versus configurations, 234
CorA, magnesium pump, 280-28 IF
Corey, Robert, 94

Cori, Gerty and Carl, 369-370, 375
Cori cycle, 360F
Cori ester, 369-3 70F

coronary heart disease and lipoprotein lipase, 507
cosubstrates, 197-199
cotranslational modifications, 690-691
coupled photosystems, 453-455T
covalent bonds, 37-38F, 120-121F, 392
citric acid cycle, 392
collagen protein structure, 120-12 IF
hydrogen bonds and, 37-38F
covalent catalysis, 169-170F
covalent modification, 158F
crassulacean acid metabolism (CAM), 471-472F
Crick, Francis H. C., 3, 573-574, 601, 635,

665, 669

Critical Assessment of Methods to Protein
Structure Prediction (CASP), 116
cyanobacteria evolution of chloroplast
photosystems, 459
cyanogen bromide (CNBr), 76-77 F
cyclic adenosine monophosphate (cAMP),
287-288F

regulatory protein activation of RNA
transcription, 653-655
cyclic electronic transfer, 452-453
cyclic guanosine monophosphate (cGMP), 287
cyclization of monosaccharide, 230-234
anomeric carbon, 23 1
anomers, 231
furanos, 23 IF

Haworth projections for, 232-234F
pyranos, 23 IF
cysteine (C, Cys), 60F
catabolism of, 540-54 IF
nomenclature, 64T
structure of, 60F
synthesis of, 523-525F
cysteine desulfurate (IscS) interactions, 11 IF
cystinuria, 544

cytidine triphosphate (CTP) synthesis,

559-560F

cytochrome bf complex, 453-455F
cytochrome b 562 , 104F
cytochrome c, 79-81F, 101F

protein structure conservation, 10 IF
sequencing, 79-8 IF

cytochrome c oxidase (electron transfer
complex IV), 431-432F
cytochromes, 221-222F
cytoplasm, 34F
cytosine (C), 8

hydrogen bonding, 38F
cytoskeleton, 20F, 23
cytosols, 20F, 23, 26F, 69 IF

D-amino acids, 57-58F
D arm, 668-669F
Dam, Henrik Carl Peter, 223
dark reactions, 443
Darwin, Charles, 15
degenerate genetic code, 667
degradation, see catabolism
dehydrogenases enzymes, 136, 203F
Delbruck, Max, 18
denaturation, 1 10-1 14F
chemical, 111-114
chaotropic agents, 111
cleavage of bonds, 112F

detergents, 111-112
disulfide bonds and bridges, 112F
double-stranded DNA, 584-585F
enzyme catalyzation, 113-114
heating, 11 IF
melting curve, 584-585F
proteins, 110-114F
renaturation and, 112-113F
deoxy sugars, 235-236F
deoxyhemoglobin, 123
deoxymyoglobin, 123
deoxyribonucleic acid, see DNA
deoxyribose, 8F, 574F

deoxythymidylate (dTMP) production, 560-564F
deoxyuridine monophosphate (dUMP)
methylation, 560-564F,
detergents, 36F

denaturation by, 112
solubility of, 36F
diabetes mellitus (DM), 381, 51 1
lipid metabolism and, 511
dialysis, 69

dichloroacetate (DCA), 408F
Dickerson, Dick, 89

dideoxynucleotides for DNA sequencing, 616, 618
dietary lipids, absorption of, 505
diffusion, 34F, 275-276
facilitated, 281
lateral, 275F

lipids in membranes, 275-276F
membrane transport and, 281
solubility and, 34F
transverse, 275-276F
diffusion- controlled reactions, 171-175
energy diagrams for, 174F
substrate binding speed and, 171-172T
superoxide dismutase, 175F
triose phosphate isomerase (TPI), 172-174F
dihydrofolate, 213F
dihydroxyacetone, 228F, 23 IF, 236F
dihydroxyacetone phosphate, 332-333F
1,25 dihydroxycholecalciferol, 218F
dipeptide, 6F, 68
diploid cells, 20
disaccharides, 236-239
cellobiose, 237-238, 239F
glucosides and, 236-239, 24 IF
glycosidic bonds in, 236-238F
lactose, 238, 239F
maltose, 237, 239F
nucleosides and, 239, 24 IF
reducing and non reducing sugars, 238-239
structures of, 237-239F
sucrose, 238, 239F

discontinuous DNA lagging strand synthesis, 608F
dissociation constant, K a, 109
acid solutions, K a , 44-48T
disulfide bonds and bridges, 1 12F
DNA (deoxyribonucleic acid), 3, 8-9F, 601-633
A-DNA, 585-586F
absorption spectrum of, 584-585F
amplification of, 615-616
bacterial, 3, 590

ball-and-stick model, 582-584F
base composition of, 579T
B-DNA, 582-584F, 586F
chromatin, 588-591
cloning vectors, 597-598F
degradation, 373
discovery of, 3
double helix, 581-585
double-stranded, 579-586
anti-parallel strands, 581-583
charge-charge interactions, 584
chemical structure of, 58 IF
complementary base pairing, 582-583F

conformations of, 585-586F
denaturation of, 5 84-5 8 5F
hydrogen bonds in, 584
hydrophobic effects, 584
major and minor grooves in, 582-583F
phosphodiester linkages (3-5') in, 580-58 IF
stability from weak forces, 583-585F
stacking interactions, 582-583F, 585T
sugar- phosphate backbones of,
van der Waal forces on, 39
ultraviolet light absorption, 584-585F
eukaryotic cells and, 20
fingerprints, 596-597F
phosphodiester linkages in, 8-9F
gene mutation, 322, 447, 469
histones, 588-590F
homologous recombination, 626-63 1
hydrogen bonds in, 37-38F
hydrolysis of, 593-596F
EcoRl and, 595-596F
nucleases and, 593-596F
restriction endonucleosis and, 593, 595T
history of, 601-602
loops for attachment of, 590, 652F
melting point, T m , 584
modified nucleotides, 564-565F
nucleic acid and, 573-574
pulling to fully extended form, 588F
recombinant, 597-598F
repair of damaged, 622-652
restriction maps, 596
sequencing of, 616-619F
single- strand, 588

space-filling model, 573F, 582-584F
sticky ends on, 598
structure of, 8-9F
supercoiled, 586-587F
synthesis, 373
Watson-Crick model, 579
Z-DNA, 586F
DNA repair, 622-625
breast cancer and, 630
excision, 624-625F

photodimerization (direct repair), 622-623
DNA replication, 602-622
base pairing in, 604-606
bidirectional, 602-603F
chromasomal, 602-603
eukaryotes, 619-622
forks, 602-603, 606, 608F, 613F
initiation (origin) of, 615F
polymerase chain reaction (PCR), 615-617F
polymerases, 603-615

chain elongation, 604-606F
interactions, 11 IF
nucleotide-group-transfer reaction,

604-605

proofreading for error correction, 607
protein types, 603-604T
replisome model, 610, 612-615
semiconservative, 602F
sequencing, 616-619F

dideoxynucleotides used for, 616, 618
parallel DNA by synthesis, 618-619
Sanger method, 616, 618
synthesis of polymerases, 607-615
binding fragments, 609-61 IF
discontinuous, 608F
Klenow fragment, 609-61 OF
lagging strands, 608-609F, 613-614F
Okazki fragments, 608-61 IF
phosphodiester linkage, 610, 612F
RNA primer for, 608-609
single-strand binding (SSB) protein, 613F
two strands simultaneously, 607-615
termination (terminus) of, 615F

774 INDEX

dnaA gene encoding, 615

Dobzhansky, Theodosius, 15

Doisy, Edward Adelbert, 223

domains, protein structure and, 101-102, 106F

Donahue, Jerry, 575

donepezil hydrochloride, 134F

double bonds, An, in fatty acids, 258-259

double helix, 581-585

anti-parallel strand formation of, 581-583
B-DNA, 582-584F

major and minor grooves in, 582-583F
stability from weak forces, 583-585F
double membranes, 273F
double-reciprocal (Fineweaver-Burk) plot,
146-147F

double-stranded DNA, 579-586
anti-parallel strands, 581-583
charge-charge interactions, 584
chemical structure of, 58 IF
complementary base pairing, 582-583F
conformations of, 585-586F
denaturation of, 5 84-5 8 5F
hydrogen bonds in, 584
hydrophobic effects, 584
major and minor grooves in, 582-583F
phosphodiester linkages (3-5') in, 580-58 IF
stability from weak forces, 583-585F
stacking interactions, 582-583F, 585T
van der Waal forces on, 39
ultraviolet light absorption, 584-585F
Drosophila melanogaster, 86, 296, 603F

E site (exit site), 682-684F
EcoRl, hydrolysis and, 595-596F
Edidin, Michael A., 276
Edman, Pehr, 74

Edman degradation procedure, 74-75F
effector enzymes, 285
eicosanoids, 268-269F
structures of, 268-269F
synthesis of, 483-486F
Eijkman, Christiaan, 198, 223
elastase, 183-185F
electrochemical cell, 317F
electrolytes, 32-34
electromotive force, 317
electron micrographs, 284, 603F
electron transfer, 319-320, 455-457
bacterial photosystems, 449-453
cyclic, 452-453
free energy, 319-320
noncyclic, 452

photosynthesis, 449-453, 455-457
Z- scheme, 455-456F
electron transport, 417-442

adenosine triphosphate (ATP) synthesis and,
417-442

chemoautotroph energy from, 439-440
cofactors, 425

enzyme complexes, 423-435

complex I (NADH to ubiquinone catalysis),
426-42 7F

complex II (succinate:ubiquinone
oxidoreductase), 427-428F
complex III (ubiquinol: cytochrome c
oxidoreductase), 428-430F
complex IV (cytochrome c oxidase),
431-432F

complex V (ATP synthase), 433-435F
Gibbs free energy change, AG, 423-425T
NADH shuttle mechanisms in eukaryotes,
436-439F

oxidation-reduction reactions, 423-425T
oxygen uptake in mitochondria, 42 IF
P/O (phosphorylated/ oxygen) ratio, 436

photosynthesis compared to, 439
protonmotive force, 421-420F
Q-cycle electron pathway, 430
reduction potentials of oxidation-reduction
components, 425T
superoxide atoms, 440-441
terminal electron acceptors and donors,
439-440

electrophiles, 39-40, 163
electrospray mass spectrometry, 72
electrostatic repulsion, 309
elongation, see chain elongation
Embden, Gustav, 331
Embden-Meyerhof-Parnas pathway, 331
enantiomers, 56

endo- envelope conformations, 234F
endocytosis, membrane transport
and, 283-284F
endonucleases, defined, 591
endoplasmic reticulum (ER), 20-2 IF, 69 IF
endosymbiotic origins, 22
energy, 10-15

activation, G*, 14F
bioenergetics, 1 1

citric acid cycle, conserved in, 405T
equilibrium and, 12-15
flow of, 1 IF

Gibbs free energy changes, 12-15
living organisms and, 10-11
metabolism, 11

NADH oxidation-reduction, conservation
from, 316-320
photosynthesis and, 1 IF
protein synthesis expense of, 684-685
reaction rates, 11-12, 14-15
thermodynamics, 12-13
energy equation, photon of light, 445, 445
energy- rich compounds, 310
enolase reactions, 338
enolpyruvate, 315F
enthalpy, H, 12

enthalpy changes, AH, 12-13, 306
Entner-Doudoroff (ED) pathway, 351-352F
entropy, S, 12

entropy change, AS, 12-13, 306
enzyme reactions, 386, 392, 394-402
aconitase, 3 96-3 9 7F

er-ketoglutarate dehydrogenase complex,
398-399F

citrate synthase, 394-396F
citric acid cycle, 386, 392, 394-402
conversion of from another, 402F
fumarase, 401

isocitrate dehydrogenase, 397-398F
malate dedrogenase, 401-402
succinate dehydrogenase complex, 399-40 IF
succinyl synthetase, 398-400F
enzyme-substrate complex (ES), 139-140,

142-143

enzymes, 2, 6-7F, 134-161, 162-195. See also
coenzymes; substrates
activation energy lowered by, 165-166F
allosteric, 153-158F

concerted (symmetry) model for, 156-157F
phosphofructokinase, 154-155F
properties of, 155-156F
regulation of enzyme activity using, 153-158
sequential model for, 157-158F
ammonia transfer from glutamate, 558
catalytic proficiency of, 144-147T
catalytic constant, k cat , 143-145
catalysts, 2, 113-114, 134
chemical reaction rates and, 15
cell cytosol behavior of, 23, 26F
citric acid cycle reactions, 386, 394-402
classes of, 136-138

oxidoreductases, 136
transferases, 136-137, 395
number system for, 137F
hydrolases, 137
lyases, 137
isomerases, 137-138
ligases, 138
co factors, 196F

conversion of from another, 402F
covalent modification of, 158F
defined, 135

electron transport, 423-435

complex I (NADH to ubiquinone catalysis),
426-42 7F

complex II (succinate:ubiquinone
oxidoreductase), 427-428F
complex III (ubiquinol: cytochrome c
oxidoreductase), 428-430F
complex IV (cytochrome c oxidase),
431-432F

complex V (ATP synthase), 433-435F
glycolysis, reactions of, 326-327T
gluconeogenesis regulation, 363-364F
inhibition, 148-153
competitive, 149-1 50F
constant, Ki,148
irreversible, 152-153F
noncompetitive, 149-151F
pharmaceutical uses of, 151-152
reversible, 148-1 52F
uncompetitive, 149-1 50F
inorganic cations and, 197
kinetic constant, k m , 144-147, 149T
kinetics and, 23, 138-149
lock-and-key theory of specificity, 180
mechanisms of, 147, 162-195
arginine kinase, 190-192F
catalysis, 166-182
cleavage reactions, 163-164
diffusion- controlled reactions, 171-175
lysozyme, 189-19 IF
nucleophilic substitution, 163
oxidation-reduction reactions, 164
serine proteases, 183-189F
transition states, 163, 164-166
metal- activated, 197
metabolite channeling, 158-159
Michaelis-Menton equation for, 140-144
multienzyme complexes, 158-159
multifunctional, 158-159
multisubstrate reactions, 147-148F
pH and rates of, 170-172F
properties of, 134-161
protein structures and, 6-7F, 113-114
reactions, 134-136F, 138-140F, 147-148
regulation of, 153-158
substrate binding and, 171-172T, 175-182F
epimers, 230

epinephrine, structure of, 63F, 199F
equilibrium, 11-15

acid dissociation constant, K a , 44-48
association constant, K a , 109-1 10F
buffered solutions, 51-52
constant, K eq , 12, 14
dissociation constant, 109
energy and, 12-15

Gibbs free energy change, AG, 12-15, 307-308
metabolic changes and, 307-308
near- equilibrium reaction, K eq , 307-308
protein-protein interactions, 109-110
rate changes and, 11-12
erythrose, 229
erythrulose, 23 IF

Escherichia coli {E. coli ), 17F, 23-24, 26F, 86F,

106, 108T

allosteric enzyme regulation and, 154-155F

Index 775

audioradiograph of replicating
chromosome, 603F

carbamoyl phosphate synthetase, 558F
cells, 17F, 23-24, 26F
chaperonin (GroE), 118-119F
covalent catalysis, 169-170F
cytochrome b 562 , 104F
flavodoxin, 105F
gloxylate pathway, 411-412
homologous recombination, 627-630
L-arabinose-binding protein, 105F
metabolic network of, 295-296
oligomeric proteins, 106, 108T
phosphofructokinase, 154-155F
ribosome, 665F, 647-675F
RNA content in, 636T
structure of, 17F, 104F
thiol- disulfide oxidoreductase, 105F
transketolase, 368F
trp operon, 688-690F
tryptophan biosynthesis enzyme, 105F
UDP N-acetylglucosamine acyl
transference, 104F
essential amino acids, 529T
essential ions, 196
ester linkages, 4-5F

ethanol, pyruvate metabolism to, 339-340F
ether, synthesis of, 487F
eukaryotes, 15-16F
chromatin and, 649
DNA replication in, 619-622
evolution and, 15-16F
glucose synthesis in, 369-370F
initiation factors, 677, 679F
mRNA processing, 656, 658-663
NADH shuttle mechanisms in, 436-439
protein synthesis and, 674-677, 679F, 691-692F
polymerases, 646-648T
ribosomes, prokaryotic cells compared to,
674-675F

RNA transcription, 646-649
secretory pathways in, 691-692F
transcription factors, 648-649T
eukaryotic cells, 18-23F
citric acid cycle and, 385
chloroplasts, 21-22F
compartmentalization, 501-502
cytoskeleton, 23
DNA and, 20

endoplasmic reticulum (ER), 20-2 IF
Golgi apparatus, 2 IF
lipid metabolism and, 501-502
metabolic pathways in, 305F
mitochondria, 21-22F
mitosis, 20F
nucleus of, 20
organelles, 19-20F
structure of, 19-20F
vesicle specialization, 22
eukaryotic DNA polymerase, 620T
eukaryotic enzymes, 364F
eukaryotic (plant) photosystems, 458-461
ATP synthase, 459-460F
chloroplasts, 458-460F
cyanobacteria evolution of, 459
organization of components, 459-460F
eukaryotic ribonucleotide reductase, allosteric
regulation of, 56 IT
eukaryotic transducers, 285
evolution, 15-17, 57-58
amino acids and, 57-58
bacterial enzymes, 364F
biochemistry and, 15-17
common ancestors, 57-58
cyanobacteria effects on chloroplast
photosystems, 459

cytochrome c sequences, 79-8 IF
endosymbiotic origins, 22
eukaryotes, 15-16F
last common ancestor (LCA), 57-58
metabolic pathways, 301-302
mitochondria and chloroplasts, 459
phylogenetic tree representation, 79-80F
prokaryotes, 15-16F
protein primary structure, 79-81
exit site (E site), 682-684F
exocytosis, membrane transport and, 283-284F
exons, 660
exonucleases, 591
extreme thermophiles, 30F

facilitated diffusion, membrane transport and, 281
fat-soluble vitamins, 198
fatty acids, 9, 257-261
anionic forms of, 258T
cis configuration, 258, 259F
coenzymes and, 215, 221
dietary requirements and, 261
double bonds, A n, in, 258-259
lipid structure of, 258-261
micromolecular structure of, 9
nomenclature, 257-258T
oxidation of, 494-501

acyl CoA synthase activation, 494
ATP generation from, 498-499
/2-oxidation, 494-50 IF
mitochondria transport, 479-498
odd- chains, 499-500
unsaturated, 500-501
polyunsaturated, 258, 260F
saturated, 258, 260F
synthesis of, 475-481, 497F
activation reactions, 479F
p - oxidation and, 49 7F
desaturation, 479-481
elongation reactions, 477-479F
extension reactions, 479-481
initiation reaction, 477F
trans configuration, 258, 259F
unsaturated, 258, 260F
feed-forward activation, 300
feedback inhibition, 300
Fenn, John B., 73
fermentation process, 340F
fibrous proteins, 86, 1 19-121. See also collagens
Filmer, David, 157
fingerprints, 77-79F, 596-597F

DNA restriction endonucleases, 596-597F
tryptic, sequencing use of, 77-79F
Fischer, Edmund (Eddy) H., 375-376
Fischer, Emil, 2, 3, 180
Fischer projections, 7F, 228-232F
aldoses, 228-230F
ketoses, 230-23 IF

monosaccharide carbohydrates, 228-232F
trioses, 228F

flavin adenine dinucleotide (FAD), 204-205F
flavin mononucleotide (FMN), 204-205F
flavodoxin, 105F
Flemming, Walter, 585
fluid mosaic model, 274-275
fluorescent protein (jellyfish), 104F
flux in metabolic pathways, 300F
FMN oxidoreductase (yeast), 105F
folate (vitamin B 9 ), 213-214F
folding, 99-103F, 114-119F
aggregation from, 119
CASP, 116

characteristics of, 1 14—1 15F
charge-charge interactions and, 117
hydrogen bonding and, 1 15-1 16F

hydrophobic effect and, 114-115
molecular chaperones and, 1 17-1 19F
pathways, 114-115F

protein stability and, 99-103F, 1 14-1 19F
tertiary protein structure and, 99-103
van der Waals interactions and, 117
forked pathways, 413F

formamidoimidazole carboxamide ribonucleotide
(FAICAR), 553F

formylglycinamide ribonucleotide (FGAR), 553F
formylglycinamidine ribonucleotide (FGAM), 553F
N-formylmethionine, structure of, 62-63F
fractional saturation, 124-125F
Franklin, Rosalind, 579
free-energy change, see Gibbs free energy
change, AG
free radicals, 164

ribonucleotide reduction, 562
freeze-fracture electron microscopy, 276-277F
fructose, 23 IF

conversion to glyceraldehyde 3 -phosphate,
348-349

gluconeogenesis regulation, 363-364F
invertase conversion to, 349
fructose 1,6 frisphosphate, 332F, 358-359F
fructose 6-phosphate, 330-33 IF, 358-359F
gluconeogenesis conversion, 358-359F
gluconeogenesis regulation, 363-364F
glycolysis conversion, 330-33 IF
Frye, L. D., 276
fuel metabolism, 295
fumarase, citrus cycle reactions, 401
fumarate, urea cycle and, 543F, 545-546F
Funk, Casimir, 198
furanos, 23 IF, 234
Furchgott, Robert F., 530

G proteins, 285-286F, 290
galactose, 229F

conversion to glucose 1 -phosphate, 349-350
galactose mutarotase, 234F
galactosides, 239, 24 IF
^aminobutyrate, structure of, 63F
gamma crystallin (cow), 104F
Gamow, George, 666
gangliosides, 265, 266F
gel-filtration chromatography, 69-70
gene, defined, 634
gene mutation, 322, 447, 469
gene orientation, 639-640F
gene regulation, 649-651, 685-690
protein synthesis, 685-690
attenuation, 688-689F
globin regulation by heme availability,
687-688F

ribosomal assembly in E. coli , 685-687F
trp operon in E. coli , 688-690F
RNA transcription and, 649-651
gene sequences, metabolism and, 295-296
genetic code, 665-668T
codons, 665-668T
degenerate, 667
history of, 665-667F
mRNA and, 666-667F
reading frames, 666-667F
tRNA and, 666, 668-670F
genetic defects, sphingolipids and, 265-266
genetically modified food, 528
genome, defined, 573
gibberellins, 270
Gibbs, Josiah Willard, 12
Gibbs free energy change, AG, 12-15, 341-342F
actual, 306, 341-342F
adenosine triphosphate (ATP), 308-312
electron transport, 423-425T

776 INDEX

enthalpy changes, AH, and, 306
entropy changes, AS, and, 306
formation of reactants, 308T
glycolysis reactions, 332, 341-342F
hydrolysis, 308-312
mass action ratio, Q, and, 306
membrane transport and, 278-279
metabolic reaction direction from, 306-312
metabolically irreversible reactions, 307,
308-312

near- equilibrium reaction, K eq , 307-308
oxidation-reduction reactions, 316-320
photosynthesis photosystems, 455-457
reduction potential and, 317-319T
standard, 306, 341-342T
thermodynamic reactions and, 12-15, 278-279
globin protein synthesis regulation, 687-688
globular proteins, 86, 122-129. See also
hemoglobin; myoglobin
gloxylate pathway, 409-412
glucokinase, 344-345F
glucolfuranose, 233F
gluconeogenesis, 303, 326F, 355-384
Cori cycle, 360F

fructose 1,6 Hsphosphate, 358-359F
glucose level maintenance (mammals), 379-381
glucose 6-phosphatase, 359-360
glucose synthesis by, 326F
glycogen metabolism, 369-372
glycogen regulation (mammals), 372-379
glycogen storage diseases, 381-382
glycolysis compared to, 356-357F
hormone regulation of, 376, 378-379F
metabolic pathway, 303
pentose phosphate pathway, 364-369
phosphoenylpyruvate carboxykinase (PEPCK)
reactions, 358F
precursors for, 360-363
acetate, 362-363
amino acids, 360-361
glycerol, 360-36 IF
lactate, 360, 361-362
propionate, 361-362
sorbitol, 362

pyruvate to glucose conversion, 356-360
pyruvate carcoxylase reaction, 357-358F
regulation of, 363-364, 376-379F
L-glucono-gamma-lactone oxidase (GULO),
210-21 IF

glucopyranose, 232F, 239F
glucose, 7-8F, 229-230F, 236F
cyclization of, 231-234F
diabetes mellitus (DM) and, 381
glycolysis, 325-354
hemeostasis phases, 380F
liver metabolic functions and, 379-380F
maintenance of levels in mammals, 379-381
monosaccharide structures of, 229-230F, 236F
pyruvate conversion via gluconeogenesis,
356-360F

pyruvate conversion via glycolysis, 3 2 8-3 2 9 F,
338-340F
solubility of, 34F
sorbitol conversion, 362G
starch and, 240-242F
storage as starch and glycogen, 240-243F
structure of, 7-8F, 34F
sugar acids derived from, 238F
sugar phosphate structures, 236F
glucose-alanine cycle, 361F
glucose 1 -phosphate, galactose conversion to,
349-350

glucose 6-phosphatase, 359-360
glucose 6-phosphate dehydrogenase deficiency, 367F
glucose 6-phosphate isomerase catalysis, 327,
330-33 IF, 345F

glucose 6-phosphate, liver metabolic functions
and, 345F

glucosides, 236-239, 24 IF
glucuronate, 238F

glutamate (E, Glu), structure of, 62F
ammonia incorporated in, 518F
catabolism of, 535

enzyme transfer of ammonia from, 558
ionization of, 65-66F
malate-aspartate shuttle, 348F
metabolic precursor use, 529
nomenclature, 64T
phosphorol group transfer, 312-313
structure of, 62F
synthesis of, 312-313, 523F
transferases catalyzation, 136-137
urea cycle and, 545-546F
phosphorol group transfer, 312-313F
glutamine (Q, Gin), structure of, 62F
ammonia incorporated in, 518F
catabolism of, 535
ligases catalyzation, 138
metabolic precursor use, 529
nomenclature, 64T
structure of, 62F
synthesis of, 312-313, 523F
glycan, 227

glyceraldehyde, 228-229F, 236F
glyceraldehyde 3-phosphate, 332-334F
fructose conversion to, 348-349
shuttle mechanisms in eukaryote, 43 7F
glyceraldehyde 3 -phosphate dehyrogenase,
333-334, 346-347F
glycerol, 360-36 IF
glyoxylate cycle, 361
gluconeogenesis precursor, 360-36 IF
oxidation of, 36 IF
glycerol 3 -phosphate, 9-1 OF

micromolecular structure of, 9-1 OF
oxidation of, 36 IF

glycerol 3 -phosphate dehyrogenase, 36 IF
glycerophospholipids, 6-1 OF, 262-265
micromolecular structure of, 9-1 OF
phosphatidates, 262-264F
plasmalogens, 263, 265F
synthesis of, 481-483F
types of, 263T

glycinamide ribonucleotide (GAR), 553F
glycine (G, Gly), 59F, 65-4T
catabolism of, 536-537F
metabolic precursor use, 529-530F
nomenclature, 64T
structure of, 59F
synthesis of, 523-524F
glycine encephalopathy, 544
glycoconjugates, 244-252
cartilage structure, 245-246F
glycoproteins, 248-252F
glycosaminoglycans, 244-245F
oligosaccharides, 248-252F
peptidoglycans, 246-248F
proteoglycans, 244-246F
glycogen, 240-243F, 369-382
cleavage of residues, 371-372F
degradation of, 371-372F, 373-374F
glucose level maintenance (mammals),
379-381

glucose storage (animals), 240-243
hormone regulation of, 376-379
linkages, 242-243F

Mendelian Inheritance in Man (MIM)
numbers, 381-382
metabolism, 369-372
molecule, 37 IF

phosphorolysis reaction, 371-372F
regulation of (mammals), 372-379, 374F

storage diseases, 381-382
synthase reaction, 370-371F
synthesis of, 369-37 IF
glycogen phosphorylase, 373-374F
degradation of, 373-375F
phosphorylated state (GPa), 375F
unphosphorylated state (GPb), 347-375F
glycolysis, 303, 325-354
aldolase cleavage, 330-332F
enolase reactions, 338

Entner-Doudoroff (ED) pathway, 351-352F
enzymatic relations of, 326-327T
fructose conversion to glyceraldehyde
3-phosphate, 348-349

galactose conversion to glucose 1 -phosphate,
349-350

Gibbs free energy change, AG, 341-342T
gluconeogenesis compared to, 356-357F
glucose catabolism, 325-354
glucose 6-phosphate isomerase catalysis, 327,
330-33 1F,345F
glucose synthesis by, 326F
glucose to pyruvate conversion by, 328-329F
glyceraldehyde 3 -phosphate dehyrogenase
catalysis, 333-334

hexokinase reactions, 326-327, 328F, 330F
history of, 331

hormone regulation of, 376, 378-379F
mannose conversion to fructose
6-phosphate, 351
metabolic pathway, 303
phosphofruktokinase- 1 (PFK-1)
catalysis, 330

phosphoglycerate kinase catalysis, 335-336
phosphoglycerate mutase catalysis, 336-337F
pyruvate kinase catalysis, 338
pyruvate metabolic functions, 338-340F
metabolism to ethanol, 339-340F
reduction to lactate, 340
regulation of, 343-347
hexokinase, 344-345
hexose transports, 343-344
metabolic pathway in mammals, 343F
Pasteur effect for, 347

phosphofruktokinase- 1 (PFK-1), 345-346F
pyruvate kinases, 346-347F
sucrose cleaved to monosaccharines, 348
triose phosphate isomerase catalysis,

332-334F

glycolytic pathway, 408

glycoproteins, 248-252F. See also oligosaccharides

glycosaminoglycans, 244-245F

glycosides, 24 IF

glycosidic bonds, 236-238F

glycosphingolipids, 256

glycosylation of proteins, 694F

glyoxylate cycle, 361

Golgi, Camillo, 2 1

Golgi apparatus, 20-2 IF, 69 IF

Goodsell, David S., 23, 34

gout, 569

Gram, Christian, 247
Gram stain, 247F
grana, 458

Greek key motif (structure), 100-101F
green filamentous bacteria, photosynthesis in,
448, 452F

Greenberg, G. Robert, 551, 552
group transfer reactions, 163
growth factors, signal transduction and, 284
guanine (G), 8, 55 IF
hydrogen bonding, 38F
structure of, 55 IF

guanosine 5'-monophosphate (GMP), 550-551F
gulose, 229F
gyrate atrophy, 544

Index 111

hairpin formation, RNA transcription, 644F

hairpin motif (structure), 100F

Haldane, J. B. S., 141

half-chair conformation, 189-190F

half-reactions, 317-319T

Haloarcula marismortui, 675, 676F

Halobacterium halobium , 270

Halobacterium salinarium , 461

Hanson, Richard, 359

haploid cells, 20

Harden, Arthur, 33 1

Haworth, Sir Walter Norman, 223, 232-234

Haworth projections, 7-8F, 232-235F

head growth, 373

heat shock proteins, 1 17-1 18F

helical wheel, 95

Helicobacter pylori, 216F

3 10 helix, 95

helix bundle motif (structure), 100F
helix-loop-helix (helix-turn-helix)
structure, 100F
heme,122-126F, 221-222F

globin protein synthesis regulation,
687-688

prosthetic groups, 122-126F, 221-222F
absorption spectra, 221-222F
cytochromes, 221-222F
hemoglobin (Hg), 122-126F
myoglobin (Mg), 122-126F
oxygen binding in, 123-126F
oxygenation and, 122
hemeostasis phases in glucose, 380F
hemiacetal, 232F
hemiketal, 232F
hemoglobin (Hb), 122-129F

allosteric protein interactions, 127-129F
a- and p - globin subunits of, 122-123F
embryonic and fetal, 126F
heme prosthetic group, 122-124F
oxygen binding, 123-129
protein structure, study of, 122-129F
protein synthesis regulation by heme
availability, 687-688
tertiary structure of, 122-123F
Henderson-Hasselbach equation, 46-47, 66
Hereditary Persistence of Fetal Hemoglobin
(HPFH), 126
Hershko, Avram, 533
heteroglycans, 240
heterotrophs, 302-303
hexokinase, glycolysis regulation of,

344-345

hexokinase reactions, 326-327, 328F, 330F
hexose transports, glycolysis regulation of,
343-344

high- density lipoproteins (HDL), 507-508
high energy bond, ~, 31 1
high-performance liquid chromatography
(HPLC), 69-70F
histamine, structure of, 63F
histidine (H, His), 61F
catabolism of, 535-536F
ionization of, 65-66F
nomenclature, 64T
structure of, 6 IF
histones, 588-590F
HIV-1 aspartic protease, 107F
Hodgkin, Dorothy Crowfoot, 88, 215, 223
Holliday, Robin, 626
Holliday junction (model) for DNA

recombination, 601, 626-627F
homocysteine, 216F
homoglycans, 240
homologous proteins, 79
homologous recombination, 626-63 1

E. coli, 627-630

Holliday junction (model), 626-627F
repair as, 631

Hopkins, Sir Frederick Gowland, 223
hopotonic cells, 35F
Hoppe-Seyler, Felix, 573
hormones, 284-287

adenylyl cyclase binding, 287-288F
G protein binding, 286
gluconeogenesis regulation by, 376,

378-379F

glycogen metabolism regulation, 376-377F
glycolysis regulation by, 376, 378-379F
lipid metabolism regulation by, 502-504
multicellular organism receptor
functions, 284-285
receptor binding, 287-288F
signal transduction and, 284-287
hydrated molecules, 34

hydrochloric acid (HCL), dissociation of, 44-45
hydrogen (H), 3, 29F

polarity of water and, 29F
hydrogen bonds, 30-32F, 37-38F
or helix, 94-97F, 98-99F
P sheets and strands, 97-99F
collagen, 120F
covalent bonds and, 37-38F
DNA (deoxyribonucleic acid), 37-38F, 584
double helix, 584
ice, formation of, 30-3 IF
interchain, 120F

loops and turns stabilized by, 98-99F
nucleic acid sites, 575-576F
orientation of, 30-3 IF
protein folding and, 115-116F
protein structures and, 94-99F
types of, 116T
water, 30-32F, 37-38F
hydrolases enzymes, 137
hydrolysis, 2, 40F, 73-74F

adenosine triphosphate (ATP), 308-312
electrostatic repulsion, 309
metabolically irreversible changes, 308-312
resonance stabilization, 310
solvation effects, 309-310
amino acid analysis and, 73-74F

chromotagraphic procedure for, 73-74F
phenylisothiocyanate (PITC) treatment, 73F
protein compositions, 74T
arsenate (arsenic) poisoning and, 336
Gibbs free energy change, AG, 308-312
nucleic acids, 591-598
alkaline, 591-592F
DNA, 593-596F
EcoRl and, 595-596F
restriction endonucleosis and, 593, 595T
ribonuclease A, 592-594
RNA, 591-594F
macromolecules, 40F
proteins, 40

signal transduction and, 285-289F
thioesters, 316
hydronium ions, 41-43
hydropathy scale, amino acids, 62T
hydrophilic substances, 32
hydrophobic effects, double-stranded DNA, 584
hydrophobic interactions, 39, 98, 114-115
hydrophobic substances, 35, 123-124F
hydrophobicity of side chains, 62
hydroxide ions, 41-43

hydroxyethylthaimine diphoshate (HETDP), 207F
hydroxyl, general formula of, 5F
hydroxylysine residue, 120F
hydroxyproline residue, 120F
hyperactivity, 359

hyperbolic binding curve, 124-126F, 146

hypertonic cells, 35F

hypoxanthine-guanine phosphoribosyl transferase
(HGBRT), 107-108F

ibuprofen, structure of, 486F
ice, formation of, 30-3 IF
idose, 229F
Ignarro, Louis J., 530
imazodole (C 3 H 4 N 2 ), titration of, 47F
immunoglobin, 129-130F
induced-fit enzymes, 179-180
inhibition, 148-153. See also regulation
antibiotics for protein synthesis, 686F
cancer drugs for, 564
competitive, 149-150F
constant, K b 148
dichloroacetate (DCA), 408F
enzyme behavior and, 148-153
kinetic constant, k m , effects on,

144-147, 149T
irreversible, 152-153F
noncompetitive, 149-15 IF
pharmaceutical uses of, 151-152, 408
phosphorylation, 687-688F
protein synthesis and, 686-688F
reversible, 148-1 52F
uncompetitive, 149-150F
inhibitors, defined, 148
initiation codons, 667, 675-679F
initiation factors, 675, 677-679F
eukaryotic cells, 677, 679F
prokaryotic cells, 677-678F
inorganic cations, 197
inosinate base pairs, 670F
inosine 5'-monophosphate (IMP) synthesis,
551-554F

inositol 1,4,5-tnsphosphate (IP 3 ), 287-289F
inositol-phospholipid signaling pathway,

287-289F

insolubility of nonpolar substances, 35-36.

See also solubility
insulin, 290-29 IF, 344F

diabetes mellitus (DM) regulation by, 381
glycogen metabolism regulation
by, 376-377F

glycolysis regulation by, 344F
receptors, 290-29 IF

integral (transmember) proteins, 270-272F
interconversions, pentose phosphate pathway,
368-369F

intermediary metabolism, 294
intermediate- density lipoproteins (IDL), 507
intermediate filaments, 23
intermediates, enzyme transition states and,
165-166F

International Union of Biochemistry and

Molecular Biology (IUBMB), 136, 401
International Union of Pure and Applied
Chemistry (IUPAC), 257
interorgan metabolism, 304-305
intrinsically disordered (unstable) proteins,
102-103

intron/extron gene organization, 660-662F
introns, 658
invertase, 349

ion- exchange chromatography, 69
ion pairing, 37
ion product, K, 42-43
ionic state of side chains, 64-65F
ionic substances, solubility of, 32-35
ionization, 41-43, 63-67
acids, 42

amino acids, 63-67
bases, 42

Henderson-Hasselbach equation for, 66

778 INDEX

ionization ( Continued )
ion product, K, 42-43
p K a values and, 63-67
titration and, 64-65F
water, 41-43

iron-sulfur clusters, 197-198F
irreversible changes, metabolic, 308-312
irreversible inhibition, 152-153F
isoacceptor tRNA molecules, 670-671
isocitrate dehydrogenase, citrus cycle reactions,
397-398F

isoleucine (I, lie), 59F, 64T
nomenclature, 64T
stereosomers of, 59F
structure of, 59F
synthesis of, 521-523F
isomerases enzymes, 137-138
isopentenyl diphosphate, cholesterol and, 488, 490
isoprenoid metabolism, cholesterol synthesis and,
490, 493-494F
isoprenoids, 256, 269F
isotonic cells, 35F

IUMBM-Nicholson metabolic chart, 504F

Jacob, Francois, 635
Johnson, W. A., 386

Karrer, Paul, 223
Kelvin scale (K), units of, 26-27
Kendrew, John C., 2-3, 88-90, 122
keto group naming convention, 399
ketohexoses, 23 IF
ketone, general formula of, 5F
ketone bodies, 508-510
lipid metabolism, 508-510
liver functions and, 509-51 OF
mitochondria oxidation and, 510
ketopentoses, 23 IF
ketoses, 228-234F

cyclization of, 230-234F
Fischer projections of, 230-23 IF
structure of, 228-230F
Khorana, H. Gobind, 666
kinases, 158, 301, 314
ATP catalyzation, 310

enzyme regulation by covalent modification
using, 158

metabolic pathway regulation and, 301
phosphorol group transfer, 314
kinetic constant, fc m , 144-147, 149T
kinetics, 23, 138-149

catalytic constant, k cat , 143-145
catalytic proficiency, 144-147T
chemical reactions, 138-139F
enzyme properties and, 138-140
enzyme reactions, 13 9- HOF
enzyme-substrate complex (ES), 139-140,
142-143

hyperbolic curve and, 146
kinetic constant, k m , 144-147, 149T
kinetic mechanisms, 147
Lineweaver-Burk (double-reciprocal) plot,
146-147F

Michaelis-Menton equation, 140-144
multisubstrate reactions, 147-148F
ping-pong reactions, 148-149F
rate (velocity) equations, 138-139, 144-145
reversible inhibitors and, 148-149T
sequential reactions, 148-149F
substrate reactions, 138-147
Klenow fragment, 609-61 OF
KNF (sequential) model for enzyme regulation,
157-158F

knob-and-stalk mitochondria structure, 433F

Knowles, Jeremy, 174
Kornberg, Arthur, 183, 601, 603, 609
Koshland, Daniel, 157
Krebs, Edwin G., 375-376
Krebs, Hans, 385-386, 397
Krebs cycle, see citric acid cycle
Kuhn, Richard, 223

L-amino acids, 57-58F
lac operon, 651-655

binding repressor to the operon, 652F
repressor blocking RNA transcription,
651-652F

repressor structure, 652-653F
cAMP regulatory protein and, 653-655F
RNA transcription activation, 653-655
lactate, 360F, 361-362
buildup, 341
Cori cycle, 360F

gluconeogenesis precursor, 360F, 361-362
oxireductases catalyzation, 136
pyruvate reduction to, 340
lactate dehydrogenase, 102F
Lactobacillus , 340
lactose, 238, 239F
lactose intolerance, 350
lagging DNA strand synthesis, 608-609F,
613-614F

Landsteiner, Karl, 250
lateral diffusion, 275F
Leloir, Luis F., 223
Lesch, Michael, 569
Lesch-Nyhan syndrome, 569
leucine (L, Leu), 59F
nomenclature, 64T
structure of, 59F
synthesis of, 521-523F
leucine zipper, 96-97A
leukotrienes, 483, 485-486F
ligases enzymes, 138
light- gathering pigments, 444-448
accessory pigments, 447-448F
chlorophylls, 444-447F
photons (energy), 445-446
resonance energy transfer, 446
special pair, 446-447F
light reactions, 443

lignin synthesis from phenylalanine, 531-532F
limit dextrins, 242
Lind, James, 209-210

Lineweaver-Burk (double-reciprocal) plot,
146-147F

linkages, 4-5F, 8-9F

micromolecular structures of, 4-5F, 8-9F
peptide bonds, 67-68F
phosphate esters, 4-5F, 8
phosphoanhydride, 4-5F, 8F
phosphodiester, 8-9F
linoleate, 48 IF

lipid anchored proteins, 272-273F
lipid metabolism, 475-513
absorption and, 505-508
dietary lipids, 505
bile salts, 505F

pancreatic lipase action, 505F
lipoproteins, 505-508F
serum albumin, 508
cholesterol, synthesis of, 488, 490-494
isoprenoid metabolism and, 490,
493-494F

level regulation, 493
steps for, 488, 490
diabetes and, 511

eicosanoids synthesis of, 483-486F
ether, synthesis of, 487F

fatty acids, synthesis of, 475-481, 49 7F
activation reactions, 479F
P~ oxidation and, 49 7F
desaturation, 479-481
elongation reactions, 477-479F
extension reactions, 479-481
initiation reaction, 477F
eukaryotic cell compartmentalization, 501-502
glycerophospholipids, synthesis of, 481-483F
hormone regulation, 502-504
IUMBM-Nicholson metabolic chart, 504F
ketone bodies, 508-510

liver functions and, 509-51 OF
mitochondria oxidation and, 510
oxidation of fatty acids, 494-501
acyl CoA synthase activation, 494
ATP generation from, 498-499
p- oxidation, 494-50 IF
mitochondria transport, 479-498
odd- chains, 499-500
unsaturated, 500-501
regulation of, 502-504
sphingolipids, synthesis of, 488-489F
triacylglycerols, synthesis of, 481-483F
lipid vitamins, 217-219F

a - tocopherol (vitamin E), 218F
cholecalciferol (vitamin D), 218-219F
phylloquinone (vitamin K), 218-219F
retinol (vitamin A), 217-218F
lipids, 9F, 256-293. See also fatty acids; lipid
metabolism; membranes
absorption of, 505-508F
anchored membrane proteins, 272-273F
bilayers, 9, 10F, 269-270, 277-278F
biological membranes, 9-1 OF, 269-270
cholesterol and, 277-278F
membrane fluidity and, 276-277
phase transition of, 277F
defined, 9

dietary absorption, 505
diffusion of, 275-276F
eicosanoids, 268-269F
fatty acids, 9, 257-261
glycerophospholipids, 262-263T
isoprenoids, 256, 269F
linkages, 4-5F

macromolecular structure of, 9F
prostaglandins, 268-269
raffs, 277

sphingolipids, 263-266F
steroids, 9, 266-268F
structural and functional diversity,

256-257F

transverse diffusion, 275-276F
triacylglycerols, 261-262F
unusual membrane compositions, 274
vesicles (liposomes), 270F, 272F
waxes, 9, 268

Lipmann, Fritz Albert, 223, 311
lipoamide, 216-217F

lipoprotein lipase, coronary heart disease and, 507
lipoproteins, 505-508F

liver metabolic functions, 344-345F, 379-380F
lock-and-key theory of specificity, 180
loop structures, a helix and p strand and sheet
connections, 98-99F
low- density lipoproteins (LDL), 507-508
lumen, 457-459F
Luria, Salvatore, 18
lyases enzymes, 137
lypoic acid, 216
lysine (K, Lys), 6 IF
catabolism, of, 542F
nomenclature, 64T
structure of, 61F
synthesis of, 520-522F

Index 779

lysosomal storage diseases, 492F
lysosomes, eukaryotic cell structure and, 20F, 22
lysozyme, 6-7, 189-191F
catalyzation by, 189-16 IF
cleavage of, 189F
conformation of, 186-190
molecular structure, 6-7F
reaction mechanism, 190-19 IF
lyxose, 229F

MacKinnon, Roderick, 280
MacLeod, Colin, 3, 573
macromolecules, 4-10
condensation of, 40-4 IF
hydrolysis of, 40F
linkages, 4-5F, 8-9F
lipids, 9

membranes, 9-10
noncovalent interaction in, 37-40F
nucleic acids, 7-9F
polysaccharides, 6-7F
proteins, 6
structure of, 4-10
magnesium (Mg), 3

major and minor grooves in double-stranded
DNA, 582-583F
malate-aspartate shuttle, 348F
malate dedrogenase, citrus cycle reactions,
401-402

malate dehydrogenase, 102F
MALDI-TOF technique, 72F
maltose, 237, 239F

mammals, metabolic pathway in, 343T
mannose, 229

conversion to fructose 6-phosphate, 351
maple syrup urine disease, 544
mass action ratio, Q, 306
mass spectrometry, 72F, 77-78F
matrix- assisted laser deabsorption ionization
(MALDI), 72

Matthaei, J. Heinrich, 337, 666
McCarty, Maclyn, 3, 573

mechanistic chemistry, 162-164. See also enzymes
melanin synthesis from tyrosine, 531, 533F
melting curve, denaturation and, 584-585F
melting point, T m , 584
membranes, 9-1 OF, 269-293
biological, 9, 269-275
chloroplasts, 458-460F
cholesterol in, 277-278F
diffusion of lipids, 275-276F
double, 273F

dynamic properties of, 275-277
fluid mosaic model of, 274-275
fluidity changes, 276-277
freeze-fracture electron microscopy, 276-277F
functions of, 269
glycerol- 3 phosphate, 9-1 OF
glycerophospholipids, 9-1 OF
lipid bilayers, 9, 10F, 269-270, 277-278F
ampithatic lipids, 270F
biological membranes, 9-1 OF, 269-270
cholesterol and, 277-278F
leaflets (monolayers) of, 270
membrane fluidity and, 276-277
phase transition of, 277F
lipid raffs, 277

lipid vesicles (liposomes), 270F, 272F
macromolecular structure of, 9-1 OF
osmotic pressure and, 34-35
photosynthesis photosystems, 457-460
plasma, 457F

protein synthesis post-translational processing
and, 691-694

oligosaccharide chains, 694F

secretory pathways, 691-692F
signal peptide, 691-692F
proteins, classes of, 10F, 270-273F
or helix, 270-271F
^barrel, 271-272F
integral (transmembrane), 270-272F
lipid anchored, 272-273F
number and variety of proteins and lipids in,
273-274F
peripheral, 272

secretions, oligosaccharides and, 252F
signal transduction across, 283-291
adenylyl cyclase signaling pathway,

287-288F

G proteins, 285-286F, 290
inositol-phospholipid signaling pathway,
287-289F

receptor tyrosine kinases, 290-29 IF
receptors, 283-285
signal transducers, 285-286
solubility and, 34-35
structure of, 10F
thylakoid, 457-460F
transport, 277-283
active, 280-283F

adenosine triphosphate (ATP), 282-283F
channels for (animal), 279-280F
characteristics of, 279T
constant, K tr , 281-282F
endocytosis and exocytosis, 283-284F
Gibbs free energy change, AG, 278-279
molecular traffic and, 277-278
passive, 280-282F
permeability coefficients, 278-279F
pores for (human), 279-280F
potential, Ai (/, 279-280F
proteins, 279-282
thermodynamics and, 278-279
menaquinone, 220F
Mendel, Gregor, 270, 447, 469
Mendelian Inheritance in Man (MIM) numbers,
381-382

Menten, Maud L., 143
Meselson, Matthew, 601
messenger RNA, see mRNA
metabolic charts, 297F
metabolic pathways, 297-302
defined, 297
evolution of, 301-302
feedback inhibition, 300
feed-forward activation, 300
flux in, 300F

forms of sequences, 297-298F

glycolysis, 325-354

glucogenesis, 354-384

regulation of, 299-301

single and multiple steps of, 298-299F

steady state in, 3 OOF

metabolic precursors, 360-363, 529-532
amino acids as, 529-532
gluconeogenesis, 360-363
metabolism, 11, 198-200T. See also glycolysis;
gluconeogenesis; metabolic pathways
adenosine triphosphate (ATP), 198-199F,

304, 308-315

allosteric enzyme phenomena, 153-154
amino acids, 514-549
amphibolic reactions, 295
anabolic (biosynthetic) reactions, 294-295F,
302-303F

autotrophs, 302-303
bacteria adaptation and, 295-296
biosynthetic (anabolic) pathways, 302303
catabolic reactions, 295F, 303-304F
cellular pathways, 302-304
citric acid cycle, 303-304

cobalamin and, 215-216F
coenzymes, 198-200T, 316-320
compartmentation, 304-305
enzyme regulation and, 153-154
experimental methods for study of, 321-322
folate (tetrahyfolate) and, 213-214
fuel, 295

gene sequences and, 295-296
Gibbs free energy change, AG, 306-312,
317-319
glucose, 303
heterotrophs, 302-303
hydrolysis, 308-312, 316
intermediary, 294
interorgan, 304-305
irreversible changes, 308-312
lipids, 475-513

nucleotide coenzymes and, 198-200
nucleotides, 550-572
nucleotidyl group transfer, 315F
oxidation and, 303-304, 316-321
phosphoryol group transfer, 312-315
reaction network of, 294-297
thioesters, 316

metabolite channeling, 158-159
metal-activated enzymes, 197
metalloenzymes, 197
methanol, 238F

methionine (M, Met), 60F, 216F

catabolism by conversion of, 539-540F
nomenclature, 64T
residue, 76

structure of, 60F, 216F
synthesis of, 520-522F
methotrexate, structure of, 550
methylation, 560-564F
cycle of reactions, 563F
deoxyuridine monophosphate (dUMP)
formation by, 560-564F
nucleotide metabolism and, 560-564F
restriction endonucleases catalysis by, 593, 595F
methylmalonyl CoA, 125-126F
Meyerhof, Otto, 331
micelles, 36F
Michaelis, Leonor, 142
Michaelis-Menton equation, 140-144
microheterogeneity, 248
microtubules, 23
Miescher, Friedrich, 573
mirror-image pairs of amino acids, 57F
Mitchell, Peter, 420
mitochondria, 21-22F, 418-421F

active transport across membrane of, 435-436
acyl CoA transport into, 497-498
adenosine triphosphate (ATP) synthesis and,
421F, 435-436
/2-oxidation and, 497-498
chemiosmotic theory, 420-423
electron transport and, 435-436
eukaryotic cell structure and, 20F, 21-22F
knob- and- stalk structure, 433F
number of, 418-419
oxidation from, 2 1
oxygen uptake in, 42 IF
photosynthesis and, 22
protonmotive force, 421-420F
pyruvate entry into, 402-405F
structure of, 419-420
mitochondrial genomes, 432F
mitosis, 20F

modified ends, mNRA, 658
molecular chaperones, 1 17—1 19F
aggregation prevention by, 1 19
chaperonin (GroE), 118-119F
heat shock proteins, 1 17-1 18F
protein folding assisted by, 1 17-1 19F

780 INDEX

molecular weight, 6

molecular weight, amino acids and, 74-75T
Monod, Jacques, 157, 635
monolayers, 36F
monosaccharides, 227-236
abbreviations for, 236T
aldoses, 228-234F
amino sugars, 235-236, 237F
ball- and- stick models of, 228F, 235F
boat conformations, 235F
chair conformations, 235F
chiral compounds, 228-230F
conformations of, 234-235F
cyclization of, 230-234
deoxy sugars, 235
derivatives of, 235-236F
endo- envelope conformations, 234F
epimers, 230

Fischer projections of, 228-232F
Haworth projections of, 232-235F
ketoses, 228-234F
sugar acids, 236, 238F
sugar alcohols, 236, 237F
sugar phosphates, 235
trioses, 226

twist conformation, 234F
monosaccharines, sucrose cleaved to, 348
Morse code, 667F

motifs (supersecondary structures), 100-101F
mRNA (messenger RNA), 9, 587, 658-663
cap formation, 658-659F
eukaryotic processing, 656, 658-663
exons, 660

genetic code and, 666-667F
intron/extron gene organization, 660-662F
introns, 658
modified ends, 658
polycistronic molecules, 679
polydenylation of, 658, 660F
protein synthesis and, 666-667F, 669-67 IF
reading frames, 666-667F
spliced precursors, 658-663
spliceosomes, 662-663F
tRNA anticodons base-paired with codons
of, 669-67 IF

wobble position, 670-671F
mucin secretions, 252F
multicellular organisms, metabolic pathways
in, 305F

multienzyme complexes, 158-159
multifunctional enzymes, 158-159
multistep pathways, 298-299F
multisubstrate enzyme reactions, 147-148F
mutagenesis, site- directed, 167, 186
Mycobacterium tuberculosis , 296
Mycoplasma pneumoniae (M. pneumoniae), 108F
myoglobin (Mb), 122-129F

heme prosthetic group, 122-123F
oxygen binding, 123-129
protein structure, study of, 122-129F
tertiary structure of, 122-123F

N-linked oligosaccharides, 249-252F
N-terminus (amino terminus), 68, 74-76F
NADH (reduced nicotinamide adenine
dinucleotide), 304, 319-320
electron transfer from, 319-320, 426-427F
glycolysis reactions, 334
metabolic reactions, 304, 319-320
shuttle mechanisms in eukaryotes, 436-439
NADPH (reduced nicotinamide adenine

dinucleotide phosphate) reduction,
466-467

Nagyrapolt, Albert von Szent-Gyorgyi, 223
near- equilibrium reaction, K eq , 307-308

negatively charged R groups, 62
Neisseria gonorrhea pilin, 105F
Nemethy, George, 157
Nephila clavipes, 121
Neurospora crassa, 212, 322
neurotransmitters, signal transduction and, 284
neutral solutions, 43
niacin (vitamin B 3 ), 200-203F
nicotinamide adenine dinucleotide (NAD), 196F,
200-203F

nicotinamide adenine dinucleotide phosphate
(NADP), 200-202F

nicotinamide mononucleotide (NMN), 200-202F

Nirenberg, Marshall, 666

nitric oxide synthesis from arginine, 530-53 IF

nitrogen (N), 3

nitrogen cycle, 515-517F

nitrogen fixation, 515

nitrogenases, 516-517

Noby, Jens G., 44

noncompetitive inhibition, 149-15 IF
noncovalent interactions, 37-40F
charge-charge, 37
hydrogen bonds, 37-38F
hydrophobic, 39-40F
ion pairing, 37
salt bridges, 37F
van der Waals forces, 38-39F
noncyclic electronic transfer, 452
nonessential amino acids, 514, 529T
nonketotic hyperglycinemia, 544
nonreducing sugars, 238-239
nonsteroid anti-inflammatory drugs
(NSAIDS), 486F
norepinephrine, 199F
nuclear magnetic resonance (NMR)
spectroscopy, 90, 321
nucleases, 591-598

alkaline hydrolysis, 591-592F
DNA, 595-596F
EcoRl and, 595-596F
endonucleases, 591
nucleic acid hydrolysis, 591-598
restriction endonucleases, 593, 595-598
ribonuclease A, 592-594
RNA, 591-593F

nucleic acids, 2, 3, 7-9F. See also DNA;
nucleosides; RNA
chromatin, 588-59 IF
cleavage of, 592F, 594F
defined, 7

double-stranded DNA, 579-586F
functions of, 573-574
history of, 573

hydrogen bond sites of, 575-576F
hydrolysis of, 591-598
alkaline, 591-592F
DNA, 593-596F
EcoRl and, 595-596F
ribonuclease A, 592-594
RNA, 591-594F
identification of, 3
macromolecular structures of, 8-9F
nucleases of, 591-598
nucleosides, 575-577F
nucleosomes, 588-590F
nucleotides as building blocks, 574-579
ribose and deoxyribose, 574F
purines and pyrimidines, 574-575F
nucleosides, 575-577F
tautomeric forms, 575-576F
restriction endonucleases, 593, 595-598
RNA in cells, 587
supercoiled DNA, 586-587F
nucleolus, 20
nucleophiles, 39-40

nucleophilic reactions, 39-41
nucleophilic substitution, 163
nucleoside triphosphates, 308-309
nucleosides, 239, 241, 575-577F
chemical structures of, 575-577F
glycosides, 239, 24 IF
nomenclature, 576-578T
nucleosomes, 588-590F
nucleotide-group-transfer reaction, 604-605
nucleotide metabolism, 550-572

adenosine 5'-monophosphate (AMP), 550-55 IF
adenosine triphosphate (ATP) reactions, 55 IF
allosteric regulation of eukaryotic
ribonucleotide reductase, 56 IT
base nomenclature, 552
cytidine triphosphate (CTP) synthesis,
559-560F

deoxythymidylate (dTMP) production, 560-564F
deoxyuridine monophosphate (dUMP)
methylation, 560-564F,

DNA and RNA modification, 564-565F
functions of, 550

guanosine 5'-monophosphate (GMP),

550- 55 IF

inosine 5'-monophosphate (IMP) synthesis,

551- 554F

5-phosphoribosyl 1 -pyrophosphate (PRPP),

55 1-552F, 555-556
purine catabolism, 565-568
purine nucleotides, synthesis of, 550-554F
purine salvage, 564-565F
pyrimidine catabolism, 568-570
pyrimidine salvage, 564-565
pyrimidine synthesis, 555-559F
ribonucleotide and deoxyribonucleotide
reduction, 560-562F
salvage pathways, 564-565
uridylate (UMP) synthesis, 556-557F
nucleotides, 198-199, 574-579
anti conformation of, 577-57 8F
chemical structure of, 574
co enzyme metabolic roles, 198-199
double-stranded DNA, 580-58 IF
nomenclature, 577-578T
nucleic acid building blocks, 574-579
nucleosides, 575-577F
purines and pyrimidines, 574-575F
ribose and deoxyribose, 574F
tautomeric forms, 575-576F
phosphodiester linkages (3-5') joining,

580-58 IF

sin conformation of, 577-578F
nucleotidyl group transfer, 315F
nucleus, eukaryotic cells, 20
Nyhan, William, 569

O-linked oligosaccharides, 249-25 IF

odd-chain fatty acids, /7-oxidation of, 499-500

Ogston, Alexander, 397

Okazaki, Reiji, 608

Okazki fragments, 608-61 IF

oligomeric protein, RNA polymerase, 363-637

oligomers (multisubunits), 103, 106, 108T

oligonucleotide- directed mutagenesis, 167

oligopeptide, 68

oligosaccharides, 227, 248-252F
ABO blood group, 250-25 IF
chain structure in post-translational
processing, 694F
diversity of chains, 248
glycosidic subclasses, 249
membrane secretions and, 252F
N-linked, 249-252F
O-linked, 249-25 IF
synthesis of, 250-251

Index 781

Online Mendelian Inheritance in Man
(OMIM), 126

organelles, eukaryotic cells, 19-20F
orotidine 5'-monophosphate (OMP), 550-55 IF
osmotic pressure, solubility and, 34-35
oxidation, 21, 164, 385, 391-394
acetyl CoA, 385, 391-394
(5- oxidation, 494-50 IF
citric acid cycle reactions, 385, 391-394
defined, 164
fatty acids, 494-501
glycerol, 36 IF

mitochondria and, 21, 497-498
oxidation-reduction reactions, 164, 200-205, 221
coenzymes, 200-205, 221, 316-320
electron transfer from, 316-320
electron transport and, 423-425T
enzyme mechanism of, 164
flavin mononucleotide (FMN), 204-205F
NADH (reduced NAD), 316-320
nicotinamide adenine dinucleotide (NAD),
200-203F

reduction potentials of electron transfer
components, 425T
thioredoxin (human), 22 IF
oxidoreductases enzymes, 136
oxygen (O), 3, 29F
sp 3 orbitals, 29F
polarity of water and, 29F
oxygen binding, 123-129
Bohr effect, 128F

allosteric protein interactions, 127-129F
carbamate adducts, 129F
conformational changes from, 124-126F
fractional saturation, 124-125F
heme prosthetic group reversibility, 123-124
hemoglobin (Hb), 123-129F
hydrophic behavior and, 123-124F
hyperbolic curve and, 124-126F
myoglobin (Mb), 123-129F
oxygenation and, 123
positive cooperativity, 124
sigmoidal (S-shaped) curves for, 124-126F
oxygen uptake in mitochondria, 42 IF
oxygenation, Calvin cycle of photosynthesis,
465-466F

oxyhemoglobin, 123
oxymyoglobin, 123

P/O (phosphorylated/ oxygen) ratio, 436

packing ratio, 588

pancreatic lipase action, 505F

papain, pH and ionization of, 170-172F

parallel jt 3 sheets, 97-98F

parallel twisted sheet, domain fold, 106F

Parnas, Jacob, 331

passive membrane transport, 280-282F
Pasteur, Louis, 2, 331

Pasteur effect for glycolysis regulation, 347
Pauling, Linus, 94
pause sites, RNA transcription, 644
Pavlov, Ivan, 183
penicillin, 247-248F
pentose phosphate pathway, 364-369
oxidative stage, 364-366F
nonoxidative stage, 364-365F, 366-368F
transketolase catalysis, 368F
interconversions, 368-369F
transaldolase catalysis, 368-369F
pepsin, 183

peptide bonds, 67-68. See also proteins
acid-catalyzed hydrolysis of, 73F
amino acids and, 67-68, 73F
hydrolysis of, 40F
peptide groups, 91-93F

cis conformation, 9 IF, 93
Ramachandran plots for, 92-93F
rotation of, 91-92F
trans conformation, 9 IF, 93
peptidyl transferase catalysis of, 681-682, 683F
polypeptide chains from, 91-93F
protein synthesis and, 681-682, 683F
residues, 67

resonance structure of, 9 IF
sequencing nomenclature, 68
structure of, 68F
peptidoglycans, 246-248F
peptidyl transferase catalysis of peptide bonds,
681-682, 683F

peptidylprolyl cis! trans isomerase (human), 104F

perchlorate (C10 4 ), 36

periodic table of elements, 4F

perioxisomes, 20F, 22

peripheral proteins, 272

permeability coefficients, 278-279F

Perutz, Max, 2-3, 88-90, 94

pH, 43-52

acid dissociation constant, K a , 44-48T

acid solutions, 43F

base solutions, 42-43F

buffered solutions, 50-52F

calculation of, 49

enzymatic rates and, 170-172F

Henderson-Hasselbach equation for, 46-47

indicators, 44F

neutral solutions, 43F

physiological uses, meter accuracy for, 44

p K a relation to 45-48T

scale, 43-44

titration of acid solutions, 47-48F
water relations to, 43T
phase transition of lipid bilayers, 277F
phenylalanine (F, Phe), 59F

lignin synthesis from, 531-532F
nomenclature, 64T
structure of, 59F
synthesis of, 524-527F
phenylanyl-tRNA, 529F
phenylisothiocyanate (PITC) treatment, 73F
amino acid treatment, 73F
Edman reagent for sequencing residues, 74-75F
phenylthiocarbamoyl (PTC) -amino acid, 73F
phosphagens, phosphoryl group transfer,

314-315F
phosphate 4-5F, 8
ester linkages, 4-5F, 8
general formula of, 5F
hydrolyses catalyzation, 137
phosphatidates, 262-264F
formation of, 48 IF

glycerophospholipid functions of, 262-264F
structure of, 264F

phosphatidylinositol 3,4,5- tnsphosphate (PIP3),
290-29 IF

phosphatidylinositol 4,5-frzsphosphate (PIP 2 ),
287-289F

5-phospho-/^D-ribosylamine (PRA), 553F
phosphoanhydride linkages, 4-5F
general structure of, 4-5F
nucleic acid structures and, 8F
phosphoarginine, 315F
phosphocreatine, 315F
phosphodiester linkages, 8-9F
DNA synthesis of, 610, 612F
nucleic acid structures and, 8-9F
nucleotides joined by (3-5') bonds,

580-58 IF

phosphoenolpyruvate (PEP), 154F, 315F, 338, 403F
phosphoenylpyruvate carboxykinase (PEPCK)
reactions, 358F, 403
phosphofructokinase, 154-155F

phosphofruktokinase- 1 (PFK-1), 330
bacterial enzyme evolution, 364F
catalysis, 330

gluconeogenesis regulation, 363-364F
glycolysis catalysis of, 330
glycolysis regulation of, 345-346F
phosphoglycerate kinase catalysis, 335-336
phosphoglycerate mutase catalysis, 3 36-3 3 7F
2-phosphoglycolate, 180-18 IF
5-phosphoribosyl 1 -pyrophosphate (PRPP),

55 1-553F, 555-556
phospholipids, 256
phosphopantetheine, 205-206F
phosphoric acid (H3PO4), titration of, 48
phosphorolysis, 371-376
glycogen reaction, 371-372F
glycogen regulation, 372-376
phosphorus (P), 3
phosphoryl, general formula of, 5F
phosphoryl group transfer, 312-315
phosphorylated state (GPa), glycogen phosphory-
lase, 375F

phosphorylation, protein synthesis regulation by,
687-688F

photoautotrophs, 303
photodimerization (direct repair), 622-623
photoheterotrophs, 303
photons (energy), 445-446
photosynthesis, 1 IF, 22, 439, 443-474
atmospheric pollution and, 457
bacterial photosystems, 448-458
coupled, 453-455T
cytochrome bf complex, 453-455F
Gibbs free energy change, AG, 455-457
internal membranes, 457
photosystem I (PSI), 448, 450-453F
photosystem II (PSII), 448-450F
reaction equations, 450T, 452T, 455T
reduction potentials, 455-457F
biochemical process, 1 IF
C 4 pathway, 469-47 IF
Calvin cycle, 443, 461-467F
carbon dioxide (C0 2 ) fixation, 461-467,
469-472

carboxysomes, 469-470F
cell structure, 22

crassulacean acid metabolism (CAM),
471-472F
dark reactions, 443
electron transport compared to, 439
energy flow, 1 IF

eukaryotic (plant) photosystems, 458-461
ATP synthase, 459-460F
chloroplasts, 458-460F
cyanobacteria evolution of, 459
organization of components, 459-460F
functions of, 443-444
light- gathering pigments, 444-448
accessory pigments, 447-448F
chlorophylls, 444-447F
photons (energy), 445-446
resonance energy transfer, 446
special pair, 446-447F
light reactions, 443
starch metabolism (plants), 467-469F
sucrose metabolism (plants), 467-469F
photosystems, 448-461
bacterial, 448-458
coupled, 453-455T
cytochrome bf complex, 453-455F
Gibbs free energy change, AG, 455-457
internal membranes, 457
photosystem I (PSI), 448, 450-453F
photosystem II (PSII), 448-450F
reaction equations, 450T, 452T, 455T
reduction potentials, 455-457F

782 INDEX

photosystems ( Continued )
eukaryotic (plant), 458-461
ATP synthase, 459-460F
chloroplasts, 458-460F
cyanobacteria evolution of, 459
organization of components, 459-460F
grana, 458
lumen, 458
stroma, 458

thylakoid membranes, 457-460F
Z- scheme, 455-456F
phycoerythrin, 447

phylloquinone (vitamin K), 218-219F
phylogenetic tree representation, 79-80F
Physeter catodon oxymyoglobin, 122F
Pinl protein, 93

ping-pong enzyme reactions, 148-149F
p K a , 45-48T, 63-67

acid dissociation parameter values, 45-48T
amino acids, ionization of and, 63-67F
buffer capacity and, 50-52F
free amino acid values, 66T
ionizable amino acid values, 168T
pFl relation to, 45-48T
titration and, 47-48F, 64-65F
plasma, lipoproteins in, 508T. See also blood
plasma

plasma membrane, 457F

plasmalogens, 263, 265F

plastoquinone, 220F

pleated J3 sheets, 97-98

polar substances, solubility of, 32-35

polarity of water, 29F

poly A tail, 658

polyacrylamide gel electrophoresis (PAGE), 70-71
polydenylation of mNRA, 658, 660F
polylinker, 597

polymerase chain reaction (PCR), 615-617F
polymerases, 603-615, 636-638

chain elongation, 604-606F, 63 7-63 8F
DNA replication and, 603-615
eukaryotic, 620T, 646T
interactions, 11 IF

nucleotide-group-transfer reaction, 604-605
proofreading for error correction, 607
protein types, 603-604T
RNA, 636-638

catalyzation by, 63 7-63 8F
chain elongation reactions, 637-638F
conformation changes, 642
eukaryotic factors, 646-648T
oligomeric protein, 363-637
transcription, 642, 646-648T
synthesis of, 607-615

binding DNA fragments, 609-61 IF
discontinuous, 608F
Klenow fragment, 609-61 OF
lagging DNA strands, 608-609F, 613-614F
Okazki fragments, 608-61 IF
phosphodiester linkage, 610, 612F
RNA primer for, 608-609
single-strand binding (SSB) protein, 613F
two DNA strands simultaneously, 607-615
polymers, 4-10

macromolecular structure of, 4-10
lipids, 9

membranes, 9-10
nucleic acids, 7-9F
proteins, 6

polysaccharides, 6-7F
polynucleotide, 7

polypeptides, 7, 68. See also proteins
polypeptide chains, 85-87, 91-93F
P strand and sheet structures, 97-99F
cotranslational modifications, 690-691
folding structures for protein stability, 99- 10 IF

peptide bonds in, 9 IF
peptide groups in, 91-93F
post-translational modifications, 690-691
protein structure from, 85-87
protein synthesis modifications, 690-69 IF
polysaccharides, 6-7F. See also carbohydrates
cellulose, 243F
chitin, 244F
glycogen, 240-243F
heteroglycans, 240
homoglycans, 240
lysozyme catalyzation of, 189-190F
micromolecular structures of, 6-7F
starch, 240-242F
structure of, 240-24 IT
polyunsaturated fatty acids, 258, 260F
pores for (human) membrane transport,

279-280F

positive cooperativity, 124
positively charged R groups, 61-62
post-transcriptional RNA modification, 655-657F
post- translational processing, 689-694
glycosylation of proteins, 694F
oligosaccharide chains, 694F
polypeptide chain modifications, 689-694F
protein synthesis, 689-694
secretory pathways, 691-692F
signal hypothesis, 691-694
signal peptide, 691-692F
signal recognition particle (SRP), 691-693F
potassium (K), 3

prenylated protein membranes, 272
primary active membrane transport, 282
primary protein structure, 67, 79-81. See also
amino acids

prochiral substrate binding, 397
prokaryotes, evolution and, 15-16F
prokaryotic cells, 17-18F
E. coli , 17F

ribosomes, eukaryotic cells compared to,
674-675F

structure of, 17-18F
proline (P, Pro), structure of, 59F
nomenclature, 64T
structure of, 59F
synthesis of, 523F

promoter recognition, RNA transcription, 641-642
promoter sequences, RNA transcription, 640-64 IF
proofreading for DNA replication error
correction, 607, 674

propionate, gluconeogenesis presursor, 361-362
prostaglandins, 268-269

lipid structure and functions, 268-269
synthesis of, 483, 485-486F
prosthetic groups, 122, 197
biotin (vitamin B 7 ), 211-212F
coenzyme behavior of, 197
cytochromes, 221-222F
defined, 122

heme, 122-126F, 221-222F
oxygen binding in, 123-126F
oxygenation and, 122
phosphopantetheine, 205-206F
pyridoxal phosphate (vitamin B 6 ), 207-209F
proteasome from yeast, 534F
Protein Data Bank (PDB), 89-90, 1 16
protein disulfide isomerase (PDI), 113-114
protein machines, 108-109F
protein synthesis, 665-696

aminoacyl-tRNA synthetases, 670-673F

antibiotic inhibition of, 686F

anticodons, 668-67 IF

codons, 665-670T, 679-684F

energy expense of, 684-685

genetic code, 665-668T

mRNA (message RNA), 666-667F, 669-67 IF

post-translational processing, 689-694
glycosylation of proteins, 694F
oligosaccharide chains, 694F
polypeptide chain modifications, 689-694
secretory pathways, 691-692F
signal hypothesis, 691-694
signal peptide, 691-692F
signal recognition particle (SRP), 691-693F
regulation of, 685-690
attenuation, 688-689F
globin, 687-688F
heme availability and, 687-688F
ribosomal assembly in E. coli , 685-687F
trp operon in E. coli , 688-690F
ribosomes, 673-68 IF, 685-687
translation, 673-684

aminoacyl-tRNA docking sites for, 680-68 IF
chain elongation, 679-684F
elongation factors, 680-68 IF
eukaryotes, initiation in, 679
initiation of, 675-679F
microcycle steps for, 679-684
peptidyl transferase catalysis, 681-682, 683F
ribosomes, 673-674
Shine-Delgarno sequence, 677F, 679
termination of, 684
translocation of ribosome, 682-684F
tRNA (transfer RNA), 665-671F, 675-681F
protein turnover, 531-533
proteins, 6-7F, 55-133
a helix, 94-97F, 98-99
allosteric, 127-129F
amino acids and, 6F, 55-84
analytical techniques, 70-74
chromatography, 73-47F
mass spectrometry, 72-73F
polyacrylamide gel electrophoresis
(PAGE), 70-7 IF

antibody binding to specific antigens,

129-130

P strands and sheets, 97-99F
biological functions of, 55-56, 119-129
classes of membrane proteins, 10F, 270-273F
coenzymes, 221

cytochrome c sequences, 79-8 IF
denaturation, 110-114F
diffusion of lipids, 275-276F
enzymes as, 6-7F
evolutionary relationships, 79-81
fibrous, 86, 119-121

folding and stability of, 99-103F, 1 14-1 19F
CASP, 116

characteristics of, 1 14—1 15F
charge-charge interactions and, 117
hydrogen bonding and, 1 15-1 16F
hydrophobic effect and, 114-115
molecular chaperones and, 1 17-1 19F
tertiary protein structure and, 99-103
van der Waals interactions and, 117
globular, 86, 122-129
glycosylation of, 694F
homologous, 79
hydrolysis of, 40F, 73-74F, 533F
linkages, 4-5F

loop and turn structures, 98-99F
macromolecular structures of, 6-7F
membranes, 10F, 270-273F
active transport, 280-283F
channels for transport (animal), 279-280F
integral (transmembrane), 270-272F
lipid anchored, 272-273F
number and variety of proteins and lipids in,
273-274F

passive transport, 280-282F
peripheral, 272

pores for transport (human), 279-280F

Index 783

oxygen binding to myoglobin and hemoglobin,
123-129

peptide bonds, 40F, 67-68F, 91-93F
phylogenetic tree representation, 79-80F
polypeptide chains, 85-87, 91-93F, 99-101F
primary structure of, 67, 79-81
protein-protein interactions, 109-1 1 1
purification techniques, 68-70
quaternary structure of, 88, 103, 106-109F
renaturation, 112-113F
secondary structure of, 87
sequencing strategies, 74-79

cleavage by cyanogen bromide (CNBr),
76-77F

Edman degradation procedure, 74-75F
human serum albumin, 78-79F
mass spectrometry, 77-78F
structure of, 85-133

binding of antibodies to antigens,

129-130F

collagen, study of, 1 19-12 IF
conformations of, 91-98, 1 10-1 14
hemoglobin (Hb), study of, 122-129F
levels of, 87-88, 99-109
loops and turns, 98-99F
methods for determining, 88-90
myoglobin (Mb), study of, 122-129F
peptide group, 91-93F
subunits, 103, 106-109F
tertiary structure of, 87F, 99-106F
ubiquitination of, 533F
UV absorbance of, 60F
proteoglycans, 244-246F
proton leaks and heat production from ATP
synthesis, 435

protonmotive force, 421-420F
proximity effect, 176-178F
psicose, 23 IF
pterin, 213-214F
purine, 8-9F, 574-575
catabolism of, 565-568
nucleotide structure, 574-575F
ring structure, 551-552F
salvage pathways, 564-565F
synthesis of nucleotides, 550-554F
nucleotides, 8-9F

puromycin, protein synthesis and, 686F
purple bacteria, photosynthesis in, 448-450F
pyranos, 23 IF, 234
pyridoxal (vitamin B 6 ), 207-209F
pyridoxal phosphate (PDP), 207-209F
pyrimidine, 8-9F, 574-575
catabolism of, 568-570
nucleotide structure, 574-575F
regulation of synthesis, 559
salvage pathways, 564-565
synthesis of, 555-559F
pyrophasphate, hydrolyses catalyzation, 137
pyrrolysine, structure of, 62-63F
pyruvate, 136-137, 315F, 338-340F, 387-391F
acetyl CoA, conversion to, 385, 387-39 IF
alanine, conversion to, 36 IF
citric acid cycle reactions, 385, 387-39 IF
gluconeogenesis conversion of, 356-360
gluconeogenesis precursor, 361
gluconeogenesis regulation, 363
glucose conversion from, 338-340F,

357-360F

glycolysis conversion of, 338-340F
lyases catalyzation, 137
metabolism to ethanol, 339-340F
mitochondria, entry into, 402-405F
oxireductases catalyzation, 136
oxidation of, 3 3 8-3 3 9F
polypeptide folding of, 101
transferases catalyzation, 136-137

pyruvate carcoxylase reaction, 357-358F
pyruvate dehydrogenase phosphorylase kinase
(PDHK), 408F

pyruvate dehydrogenase structural core, 108F
pyruvate kinase, 101, 338, 346-347F
glycolysis catalysis of, 338
glycolysis regulation of, 346-347F
reduction to lactate, 340

Q- cycle electron pathway, 430
quaternary protein structure, 88, 103, 106-109F
Escherichia coli (E. coli ) oligomeric
proteins, 108T
examples of, 107F

oligomers (multisubunits), 103, 106, 108T
protein machines, 108-109F
subunits, 103, 106-109F

R group amino acids, see side chains
R (relaxed) state, 126
racemization, 58
Racker, Efriam, 461
Ramachandran plots, 92-93F
Ramachandran, G. N., 92, 1 19
rate (velocity) equations, 138-139, 144-145
reaction coordinates, 165-166F
reactions, metabolic network of, 294-297F
reactive center, 196
reading frames, 666-667F
receptors, 283-285
recombinant DNA, 597-598F
recombination, see homologous recombination
reduced nicotinamide adenine dinucleotide,
see NADH

reducing sugars, 238-239
reduction, 164. See also oxidation-reduction
Calvin cycle of photosynthesis, 466-467
defined, 164

deoxyribonucleotide, 560-562F
ribonucleotide, 560-562F
reduction potential, 317-319T, 425T, 455-457T
coenzymes, 317-319T

electron transport oxidation-reduction compo-
nents, 425T

photosynthesis, 455-457F
reductive pentose phosphate cycle, see Calvin cycle
regeneration, Calvin cycle of photosynthesis,
466-467F

regulation, 153-158, 343-347, 363-364. See also
inhibition

citric acid cycle, 406-407F
enzyme activity, 153-158

allosteric enzymes, 153-158F
concerted (symmetry) model for, 156-157F
cooperative binding and, 156F
covalent modification, 158F
phosphofructokinase, 154-155F
sequential (KNF) model for, 157-158F
sigmoidal (S shaped) curves for, 153F, 156F
gluconeogenesis, 363-364F
glycolysis, 343-347
hexokinase, 344-345
hexose transports, 343-344
metabolic pathway in mammals, 343F
Pasteur effect for, 347

phosphofruktokinase- 1 (PFK-1), 345-346F
pyruvate kinases, 346-347F
hormones for, 502-504
IUMBM-Nicholson metabolic chart, 504F
lipid metabolism, 502-504
protein synthesis, 685-690
attenuation, 688-689F
globin, 687-688F
heme availability and, 687-688F

ribosomal assembly in E. coli , 685-687F
trp operon in E. coli , 688-690F
relative molecular mass, 6
renal glutamine metabolism, 547-548
renaturation, 112-113F
replisome, defined, 603
replisome model, 610, 612-615
residues, 5, 67-68F, 74-75F

amino acids, 67-68F, 74-75F, 166-168T
(3 strand and sheet turns, 99F
catalysis and, 166-168T
catalytic frequency distribution, 168T
collagen and formation of, 120-12 IF
Edman degradation procedure for, 74-75F
glycogen, cleavage of, 371-372F
ionizable amino acid functions, 166-168T
macromolecule structure of, 5
methionine, 76

peptide bond linkages, 67-68F
phenylisothiocyanate (PITC) treatment,
74-75F

p K a values of ionizable amino acids, 168T
protein structure and, 120-12 IF
sequences of, 68, 74-75F
resonance energy transfer, 446
resonance stabilization, 310
respiration process, 340
restriction endonucleases, 593, 595-598
defined, 593
DNA and, 593,595-598
DNA fingerprints, 596-597F
hydrolysis and, 593,595
methylation, 593, 595F
nucleic acids and, 593, 595-598
recombinant DNA, 597-598F
restriction maps, 596
specificities of, 595T
types I and II, 593, 595
restriction maps, 596
retinol (vitamin A), 217-218F
retinol-binding protein (pig), 104F
reverse turns, protein structures, 99
reversible inhibition, 148-1 52F
rho- dependent RNA transcription termination,
644-645F

Rhodopseudomonas photosystem, 107F
Rho do spirillum rubrum , 484
riboflavin, 204-205F
ribofuranose, 233F

ribonuclease A (Rnase A), 90F, 1 1 1-1 13F
denaturation and renaturation
of, 112-113F
disulfide bridges in, 1 12F
heat denaturation of, 1 1 IF
hydrolysis by, 592-594
ribonucleic acid, see RNA
ribopyranose, 233F
ribose, 7, 229F, 236F, 574F
cyclization of, 232-233F
monosaccharide structures of, 229F, 236F
nucleotide structure, 574F
sugar phosphate structure, 236F
ribosomal RNA, see rRNA
ribosomes, 108F, 673-681F

aminoacyl-tRNA binding sites in, 675, 677F
chain elongation and, 673-674, 682-684F
eukaryotic versus prokaryotic cells,
interactions, 11 IF

protein synthesis, 673-68 IF, 685-687
regulation of protein synthesis, 685-687F
rRNA composition of, 674-675F
translocation by one codon, 682-684F
ribulose, 230-23 IF
ribulose 1,5-Hsphosphate, 465-466F
ribulose 5-phosphate conversion, 367F
right turn structures, 98-99F

784 INDEX

RNA (ribonucleic acid), 3, 9, 634-664
cell content, 587
classes of, 587
cleavage, 594F, 655-657F
discovery of, 3

eukaryotic mRNA processing, 656, 658-663
hydrolysis, 591-594
alkaline, 591-592F
nucleases and, 591-594
ribonuclease A, 592-594F
lac operon, 651-655

binding repressor to the operon, 652F
cAMP regulatory protein and, 653-655F
repressor blocking transcription, 651-652F
repressor structure, 652-653F
transcription activation, 653-655
messenger (mRNA), 9, 587, 656, 658-663
modified nucleotides, 564-565F
molecule types, 9
polymerase, 108F, 11 IF, 636-638
catalyzation by, 63 8F
chain elongation reactions, 63 7-63 8F
interactions, 11 IF
multisubunit, 108F
oligomeric protein, 363-637
post-transcriptional modification of, 655-657
ribosomal (rRNA) processing, 656-657F
transfer (tRNA) processing, 655-657F
ribosomal (rRNA), 9, 587, 656-657F
small nuclear (sRNA), 662-663F
stem-loop structures, 587-588F
synthesis of, see transcription
transfer (tRNA), 9, 587, 655-657 F
types of, 635-636
RNA polymerase, 108F, 1 1 IF
RNA primer for DNA synthesis, 608-609
RNA transcription, 639-651

cAMP regulatory protein activation of, 653-655
eukaryotes, 646-649
chromatin and, 649
polymerase reactions, 646-648T
transcription factors, 648-649T
gene regulation, 649-651
initiation, 639-643
or subunits, 641-642T
gene orientation, 639-640F
polymerase changes in conformation, 642
process of, 643F
promoter recognition, 641-642
promoter sequences, 640-64 IF
lac repressor blockage of, 651-652F
termination, 644-645
hairpin formation, 644F
pause sites, 644
rho- dependent, 644-645F
rofecoxib (Vioxx), structure of, 486F
Rose, Irwin, 533

rRNA (ribosomal RNA), 9, 587, 656-657
cleavage, 656-65 7F

post-transcriptional modification, 656-657 F
protein synthesis and, 674-675F
ribosome composition of, 674-675F
RS amino acid system configuration, 6 IF
rubisco (rubilose 1,5-Hsphosphate carboxylase-
oxygenase), 462, 464-466F

S-adenosylmethionine, 199F

saccharides, see carbohydrates; polysaccharides

Saccharomyces cerevisiae, 296F

salicylates, 486

Salmonella typhymurium , 514F, 528F

salt bridges, 37F

salvage pathways, 564-565

Sanger method for DNA sequencing, 616, 618

Sanger, Frederick, 616

saturated fatty acids, 258, 260F
Schiff bases, 121F, 208F, 332-333F
scurvy, ascorbic acid and, 209-210
seawater, properties of, 33F
second messengers, 285

secondary active membrane transport, 282, 283F
secondary protein structure, 87
secretory pathways, 691-692F
selenocysteine, structure of, 62-63F
semiconservative DNA replication, 602F
semiquinone anion, 220F
sequencing, 68, 74-81, 616-619
amino acid residues, 68, 74-75F
C-terminus (carboxyl terminus), 68, 76F
cytochrome c, 79-8 IF
DNA, 77F, 616-619F

dideoxynucleotides used for, 616, 618
parallel strands by synthesis, 618-619
Sanger method, 616, 618
Edman degradation procedure for, 74-77F
evolution relationships and, 79-8 IF
human serum albumin, 78-79F
N-terminus (amino terminus), 68, 74-76F
protein strategies, 76-79F

cleavage by cyanogen bromide (CNBr),
76-77 F

human serum albumin, 78-79F
mass spectrometry, 77-78F
tryptic fingerprint, 77-79F
sequential enzyme reactions, 148-149F
sequential (KNF) model for enzyme regulation,
157-158F

serine (S, Ser), 56-57F, 60-61F
catabolism of, 536-537F
metabolic precursor use, 529-530F
nomenclature, 64T

RS amino acid system configuration, 6 IF
structure of, 56-57F, 60-6 IF
synthesis of, 523-524F
serine proteases, 183-189F
catalytic triad, 185F
catalysis modes for, 185-188
chymotrypsin, 183-188F
elastase, 183-185F
substrate binding, 186-188F
substrate specificity of, 184-185
trypsin, 183-185F

zymogens as inactive enzyme precursors,
183-184

serum albumin (human), 78-79F, 104F, 508
Shine-Delgarno sequence, 677F, 679
shuttle mechanisms, 436-439F
malate-aspartate shuttle, 348F
NADH in eukaryotes, 436-439F
shuttle mechanisms in eukaryote, 43 7F
side chains, 56, 59-62

alcohol groups with, 60-61
aliphatic R groups, 59
a helix proteins, 95
amino acid structure and, 56, 59
aromatic R groups, 59-60
hydrophic effect on, 114-115
hydrophobicity of amino acids with, 62
ionic states of, 64-65F
negatively charged R groups, 62
positively charged R groups, 61-62
protein folding and, 115-116
sulfur- containing R groups, 60
sigmoidal (S-shaped) curves, 124-126F, 153F, 156F
signal hypothesis, 691-694
signal peptide, 691-692F
signal recognition particle (SRP), 691-693F
signal transduction, 283-291

adenylyl cyclase signaling pathway, 287F

G proteins, 285-286F, 290

hormones receptors and binding, 284-287

hydrolysis and, 285-289F
inositol-phospholipid signaling pathway,
287-289F

insulin receptors, 290-29 IF
membrane cells, 283-291
pathways, 284-285, 287-289F
receptor tyrosine kinases, 285, 290-29 IF
receptors, 283-285
transducers, 285-286

sin conformation of nucleotides, 577-578F

single step pathways, 298-299F

single-strand binding (SSB) protein, 613F

single-strand DNA, 588

site-directed mutagenesis, 167, 186

small nuclear ribonucleic acid (snRNA), 662-663F

Smith, Michael, 167

sn-glycerol 3-phoshphate, 484

Soderbaum, H. G., 196

sodium (Na), 3

sodium chloride (NaCl), 33F, 37
sodium dodecyl sulfate (SDS), 36F
sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE), 71F
sodium palmitate, 36
solubility, 32-36

amphipathic molecules, 36
cellular concentrations, 34F
chaotropes, 36
detergents, 36F
diffusion, 34F
electrolytes, 32-34
hydrated molecules, 34
hydrophilic substances, 32
hydrophobic substances, 35
ionic and polar substances, 32-35
nonpolar substances, 35-36
osmotic pressure, 34-35
solvated molecules, 34
surfactants, 36
water and, 32-36
solubilization, 36
solvated molecules, 34
solvation effects, 309-310
sorbitol conversion from glucose, 362G
sorbose, 23 IF

Sorensen, Soren Peter Lauritz, 44
sp 3 orbitals, 29F
space-filling models, 90F
DNA, 573F, 582-584F
proteins, 90F
special pair, 446-447F
specific heat of water, 3 1
sphingolipids, 263-266F
cerebrosides, 265, 266F
ceremide, 264, 265F
gangliosides, 265, 266F
genetic defects and, 265-266
pathways for formation and degradation
of, 492F

sphingomyelins, 264, 265F
synthesis of, 488-489F
sphingomyelins, 264, 265F
spider silk strength, 121
spindle fibers, 56

spliced precursors, mNRA, 658-663
spliceosomes, 662-663F
squalene, cholesterol and, 488, 490
stacking interactions, double-stranded DNA,
582-583F, 585T
Stahl, Franklin, 601

Staphylococcus aureus (S. Aureus), 76, 247-248F
starch, 240-243
amylase, 242F
amylose, 24 IF
amylopectin, 241-242F
digestion of, 241-242

Index 785

glucose storage (plants), 240-243F
metabolism (plants), 467-469F
structure of, 240-24 IT
synthesis of, 467-468F
starch, 240-243, 467-469
steady state, metabolic pathways, 300F
steady-state derivation, 141-142
stem length mutation, 270
stem-loop structures in RN, 587-588F
stereochemical numbering, 484
stereoisomers, 56, 59F
stereospecifity, 134-135
steroids, 9, 266-268F

cholesterol and, 266-268
isoprene structure of, 266F
lipid structures of, 266-267F
micromolecular structure of, 9
signal transduction and, 285
Strandberg, Bror, 89
Streptococcus pneumoniae , 3
Streptomyces, potassium channel protein, 107F
stroma, 458

substrates, 90F, 134-148, 175-182

binding properties, 139-140, 176, 178-181F,
185-188

binding sites, 90F, 674F
binding speed, 171-172T
diffusion- controlled reactions, 171-172T
enzymatic catalysis modes and, 175-182
induced fit, 179-180
proximity effect, 176-178F
transition-state stabilization, 176,
180-182F

weak binding and, 176, 179— 18 IF
enzyme kinetics and, 138-148
enzyme reactions, 134-135, 138-147
enzyme-substrate complex (ES), 139-140,
142-143

Michaelis-Menton equation for, 140-144
multisubstrate reactions, 147-148F
prochiral binding, 397
rate (velocity) equations for, 138-139F,
144-145F

serine proteases and, 186-188F
specificity of, 184-185
stereospecifity of, 134-135
subunits, 103, 106-109F
succinate dehydrogenase complex, citrus cycle
reactions, 399-40 IF

succinate:ubiquinone oxidoreductase (electron
transfer complex II), 427-428F
Succinyl Co A, 216F

catalyzed structure of, 216F
thioester hydrolysis, 316
succinyl synthetase, citrus cycle reactions,
398-400F
sucralose, 240
sucrose, 238-239F

cleaved to monosaccharines, 348
metabolism (plants), 467-469F
structure of, 238-239F
synthesis of, 467-469F
sugar acids, 236, 238F
sugar alcohols, 236, 237F
sugar phosphates, 235
sugars, 235-236, 238-239
abbreviations for, 236T
disaccharides, 238-239
monosaccharides, 235-236F
nonreducing, 238-239
reducing, 238-239
sulfhydryl, general formula of, 5F
sulfur (S), 3

sulfur- containing R groups, 60
Sumner, James B., 135
supercoiled DNA, 586-587F

superoxide anions, 440-441

superoxide atoms, 440-441

superoxide dismutase, 175F

supersecondary structures (motifs), 100-101F

surfactants, solubility of, 36

sweetness receptors, 240

symport, membrane transport, 280-28 IF

Synechococcus elongatus , 470F

synonymous codons, 667

synthase, 395

ATP catalysis, 43 3-43 5F
defined, 395

glycogen reaction, 370-37 IF
synthesis, 13

adenosine triphosphate (ATP), 417-442
amino acids, 520-529
cancer drug inhibition of, 564
defined, 13

DNA, two strands simultaneously, 607-615
nucleotide metabolism and, 550-559
proteins, 665-696
purine nucleotides, 550-554F
pyrimidine, 555-559F
synthetase, defined, 395
Systeme International (SI) units, 26-2 7T

T (tense) state, 126

tagatose, 23 IF

tail growth, 373

talose, 229F

Tanaka, Koichi, 73

Tatum, Edward, 212, 634

tautomeric forms of nucleic acids, 575-576F

terminal electron acceptors and donors, 439-440

termination (stop) codons, 667F, 682, 684

terpenes, 256

tertiary protein structure, 87F, 99-106F
cytochrome c structure conservation, 10 IF
domains, 101-102, 106F
examples of, 104-105F
hemoglobin (Hb), 122-123F
intrinsically disordered (unstable) proteins,
102-103

motifs (supersecondary structures), 100-101F
myoglobin (Mb), 122-123F
polypeptide folding and stability of, 99-10 IF
protein stability and, 99-103
supersecondary structures (motifs), 100-101F
tetrahydro folate, 213-214F
thermodynamics, 12-15, 278-280
activation energy, Gf, 14F
equilibrium constant, K eq , 12, 14
Gibbs free energy change, AG, 12-15,

278-279

membrane potential, Ai )/, 279-280F
membrane transport and, 278-280
reaction rates and, 14-15
Thermus thermophilius , 675, 676F
thiamine (vitamin BJ, 206-207F
thiamine diphosphate (TDP), 206-207F
thiamine pyrophosphate (TPP), 206
Thiobacillus , 303F
thiocyanate (SCN), 36
thioesters, hydrolysis of, 316
thiol (sulfhydryl), general formula of, 5F
thiol- disulfide oxidoreductase, 105F
thioredoxin (human), 105F

coenzyme oxidation-reduction, 22 IF
oxidized, 22 IF
structure of, 105F
threonine (T, Thr), 58, 60-61F
catabolism of, 537-538
nomenclature, 64T
structure of, 58, 60-6 IF
synthesis of, 520-522F

threose, 229

thylakoid membranes, 457-460F
thymine (T), 8-9F
thyroxine, structure of, 63F
titration, 47-48F

acetic acid (CH 3 COOH), 47F
acid solutions, 47-48F
amino acids, 64-65F
imazodole (C 3 H 4 N 2 ), 47F
ionization and, 64-65F
phosphoric acid (H3PO4), 48
pK a values from, 45-48T, 64-65F
T y/C arm, 668-669F
trans conformation, 9 IF, 93, 258, 259F
transaldolase catalysis, 368-369F
transanimation reactions, ammonia assimilation
and, 518-519F
transducers, 285-286
bacterial, 285-286
eukaryotic, 285
G proteins, 285-286F

membrane signal transduction and, 285-286
transduction, see signal transduction
transfer RNA, see tRNA
transferases enzymes, 136-137, 395
transition-state stabilization, 180-182F
transition states, 163, 164-166
activation energy, 165F
catalyst stabilization for, 164-166
defined, 163

enzyme mechanisms and, 164-166
intermediates and, 165-166F
nucleophilic substitution, 163
reaction coordinates, 165-166F
transketolase catalysis, 368F
translation, 673-684. See also post-translational
processing

chain elongation, 679-684F
aminoacyl-tRNA docking sites
for, 680-68 IF

elongation factors, 680-68 IF
microcycle steps for, 679-684
peptidyl transferase catalysis, 681-682F
translocation of ribosome, 682, 684F
initiation of, 675-679F
eukaryotes, 679

initiation factors, 675, 677-679
ribosomes, 673-674
Shine-Delgarno sequence, 677F, 679
tRNA initiator, 675, 677F
protein synthesis and, 673-684
ribosomes and, 673-675F, 677F
aminoacyl-tRNA binding sites
for, 675, 677F

eukaryotic versus prokaryotic, 674-675F
subunit composition of, 674-675F
Shine-Delgarno sequence, 675F, 679
termination of, 684

transmember (integral) proteins, 270-272F
transport, see electron transport; membranes
transport constant, K tr , 281-282F
transverse (flip-flop) diffusion, 275-276F
triacylglycerols, 261-262F
digestion of, 262
structure of, 26 IF
synthesis of, 481-483F
Trichodesmium, 515F
triene, defined, 486

trifunctional enzymes, /7-oxidation and, 498
triiodothryonine, structure of, 63F
triose phosphate isomerase (TPI), 107F,

172-174F

catalysis, 332-334F

diffusion- controlled reactions, 162F, 172-174F
trioses, 226
tripeptide, 68

786 INDEX

tRNA (transfer RNA), 9, 587, 655-657, 665-671,
675-681

aminoacyl-tRNA synthetases, 670-673F
anticodons, 668-67 IF
cleavage, 655-656F
base-pairing, 669-670F
cloverleaf structure, 668-669F
genetic code and, 669-670F
isoacceptor molecules, 670-671
mRNA codons base-paired with anticodons
of, 669-670F

post-transcriptional modification, 655-657F
protein synthesis and, 665-67 IF, 675-68 IF
three-dimensional (tertiary) structure of,
668-669F, 680

translation initiator, 675-68 IF
Watson-Crick base pairing, 670F
wobble position, 670-671F
trp operon, protein synthesis regulation by, 688-690F
trypsin, 76-77F, 183-185F
tryptic fingerprint, sequencing and, 77-79F
tryptophan (W, Trp), 58-60F
nomenclature, 64T
structure of, 58-60F
synthesis of, 524-52 7F
tryptophan biosynthesis enzyme, 105F
turn structures, a helix and strand and sheet
connections, 99F
twist conformations, 234F
type III triple helix, 1 19F
tyrosine (Y, Tyr), 58-60F
catabolism of, 541-542F
melanin synthesis from, 531, 533F
nomenclature, 64T
structure of, 58-60F
synthesis of, 524-527F

ubiquinol, 220

ubiquinokcytochrome c oxidoreductase (electron
transfer complex III), 428-430F
ubiquinone (coenzyme Q), 2 19-22 IF
ubiquitin, 533F

ubiquitination of proteins, 533F
UDP N-acetylglucosamine acyl transference, 104F
ultraviolet light absorption in double-stranded
DNA, 584-585F

uncompetitive inhibition, 149-1 50F
uncouplers, 420-42 IF
uniport, membrane transport, 280, 28 IF
units for biochemistry, 26-27T
unphosphorylated state (GPb), glycogen
phosphorylase, 347-375F
unsaturated fatty acids, 258, 260F, 500-501
uracil (U), 8

urea, structure of, 1 12
urea cycle, 542-547

amino acid metabolism and, 542-547
ancillary reactions to, 547
carbamoyl phosphate synthesis, 543 F
conversion of ammonia to urea, 542-547
reactions of, 543-546F
uric acid, 566-569F

uridine diphosphate glucose (UDP-glucose),
200-20 IF

uridine triphosphate (UTP), 200-20 IF
uridylate (UMP) synthesis, 556-557F
UV absorbance of proteins, 60F

vacuoles, 20F, 22
valine (V, Val), 59F
nomenclature, 64T
structure of, 59F
synthesis of, 521-523F
van der Waals, Johannes Diderik, 38
van der Waals forces, 38-39F
van der Waals interactions, 117
van der Waals radii, 39T
vaporization of water, 32
variable arm, 668-669F
vesicles, 20F, 272F
eukaryotic cells, 22
liposomes, 270F, 272F
specialization, 20F
vitamins, 196, 198-199T

ascorbic acid (vitamin C), 209-211
biotin (vitamin B 7 ), 21 1-21 2F
cobalamin (vitamin B 12 ), 215-216F
deficiencies, 198T, 209-210, 214, 215
fat-soluble, 198
folate (vitamin B 9 ), 213-214F
functions of, 197-199T
history of, 198
lipid, 21 7-2 19F

or- tocopherol (vitamin E), 218F
cholecalciferol (vitamin D), 218-219F
phylloquinone (vitamin K), 218-219F
retinol (vitamin A), 217-218F
niacin (vitamin B 3 ), 200-203F
pyridoxal (vitamin B 6 ), 207-209F
sources, 199T

thiamine (vitamin B^, 206-207F
water-soluble, 198
Voss-Andreae, Julian, 127

Walker, John E., 223
Warburg, Otto, 386
warfarin (rat poison), 220F

water, 28-54

acid disolution constants, 44-48
buffered solutions, 50-52
chemical properties of, 28, 39-52
concentration of, 41F
condensation of, 40-4 IF
hydrogen bonding in, 30-32, 37-38F
ice, formation of, 30-3 IF
insolubility of nonpolar substances, 35-36
ionization of, 41-43T
noncovalent interactions, 37-40F
charge-charge, 37
hydrogen bonds, 37-38F
hydrophobic, 39-40F
van der Waals forces, 38-39F
nucleophilic reactions, 39-41
pH scale and, 43-44, 49-52
physical properties of, 28-39
polarity of, 29F

solubility of ionic and polar substances, 32-35
specific heat of, 31
vaporization of, 32
water-soluble vitamins, 198
Watson, James D., 3, 573-574, 575, 601
Watson-Crick base pairing, 668-670F
Watson-Crick DNA model, 579, 601
waxes, lipid structure and functions, 9, 268
weak substrate binding, 179-179F
website accuracy, 401
Wilkins, Maurice, 579
Williams, Ronald, 420
Windaus, Adolf Otto Reinhold, 223
wobble position, 670-67 IF
Wohler, Friedrich, 2
Wyman, Jeffries, 157

X-ray crystallography, 88-90F
X-ray diffraction pattern, 88F
xylose, 229F
xylulose, 23 IF

yeast, 105F, 345-347F

FMN oxidoreductase, 105F
octamer enzyme, 345-346
proteasome from, 534F
pyruvate kinase regulation by, 347F
Young, William John, 331

Z-DNA, 586F

Z- scheme, photosynthesis path, 455-456F
zwitterions (dipolar ions), 56
zymogens, 183-184

Common Abbreviations in Biochemistry

ACP

ADP

AMP

cAMP

ATP

1,3BPG

2,3BPG

CDP

CMP

CoA

CTP

DHAP

DNA

cDNA

DNase

E°

E of

emf

ETF

FAD

FADH 2

F1,6BP

FMN

fmnh 2

F6P

AG°'

GDP

GMP

cGMP

G3P

G6P

GTP

HDL

HETPP

HPLC

IDL

elF

IMP

IP 3

^•cat

K e q

K m

FDF

FHC

M r

acyl carrier protein

adenosine 5 '-diphosphate

adenosine 5 '-monophosphate (adenylate)

3 ',5 '-cyclic adenosine monophosphate

adenosine 5 '-triphosphate

base pair

1,3- fcphosphoglycerate

2,3 - fcphosphoglycerate

cytidine 5 '-diphosphate

cytidine 5 '-monophosphate (cytidylate)

coenzyme A

cytidine 5 ' -triphosphate

dihydroxyacetone phosphate

deoxyribonucleic acid

complementary DNA

deoxyribonuclease

reduction potential

standard reduction potential

elongation factor

electromotive force

electron-transferring flavoprotein

Faraday’s constant

flavin adenine dinucleotide

flavin adenine dinucleotide (reduced form)

fructose 1,6-frzsphosphate

flavin mononucleotide

flavin mononucleotide (reduced form)

fructose 6-phosphate

actual free-energy change

standard free-energy change

guanosine 5 '-diphosphate

guanosine 5 '-monophosphate (guanylate)

3 ',5 '-cyclic guanosine monophosphate

glyceraldehyde 3 -phosphate

glucose 6-phosphate

guanosine 5 '-triphosphate

enthalpy

hemoglobin

high density lipoprotein

hydroxyethylthiamine pyrophosphate

high-pressure liquid chromatography

intermediate density lipoprotein

initiation factor

eukaryotic initiation factor

inosine 5 '-monophosphate

inositol 1,4,5-trzsphosphate

acid dissociation constant

catalytic constant

equilibrium constant

Michaelis constant

kilobase pair

low density lipoprotein

light-harvesting complex

relative molecular mass

myoglobin

NAD©

NADH

NADP©

NADPH

NMN©

NDP

NMP

NTP

dNTP

PAGE

PCR

2PG

3PG

PEP

PFK

PIP 2

PEP

PPi

PQH 2

PRPP

PSI

PSII

QH 2

RNA

mRNA

rRNA

snRNA

tRNA

RNase

snRNP

RPP

Rubisco

dTDP

dTMP

TPP

dTTP

UDP

UMP

UTP

v 0

VFDF

XMP

nicotinamide adenine dinucleotide
nicotinamide adenine dinucleotide (reduced form)
nicotinamide adenine dinucleotide phosphate
nicotinamide adenine dinucleotide phosphate
(reduced form)

nicotinamide mononucleotide
nucleoside 5 '-diphosphate
nucleoside 5 '-monophosphate
nucleoside 5 '-triphosphate
deoxynucleoside triphosphate
inorganic phosphate (or orthophosphate)
polyacrylamide gel electrophoresis
polymerase chain reaction
2-phosphoglycerate
3 -phosphoglycerate
phosphoenolpyruvate
phosphofmctokinase
isoelectric point

phosphatidylinositol 4,5-frzsphosphate

pyridoxal phosphate

inorganic pyrophosphate

plastoquinone

plastoquinol

5-phosphoribosyl 1 -pyrophosphate

photo system I

photosystem II

ubiquinone

ubiquinol

release factor

ribonucleic acid

messenger ribonucleic acid

ribosomal ribonucleic acid

small nuclear ribonucleic acid

transfer ribonucleic acid

ribonuclease

small nuclear ribonucleoprotein

reductive pentose phosphate

ribulose 1,5-frzsphosphate carboxylase-oxygenase

entropy

deoxythymidine 5 '-diphosphate
transcription factor

deoxythymidine 5 '-monophosphate (thymidylate)

thiamine pyrophosphate

deoxythymidine 5 '-triphosphate

uridine 5 '-diphosphate

uridine 5 '-monophosphate (uridylate)

uridine 5 '-triphosphate

velocity

maximum velocity

initial velocity

very low density lipoprotein

xanthosine 5 '-monophosphate

Abbreviations for amino acids are given on pages 57-62, and those
for major pyrimidine and purine bases are given on page 575.

First position
(5' end)

Second position
C A

Third position
(3' end)

Phe

Ser

Tyr

Cys

Phe

Ser

Tyr

Cys

Leu

Ser

STOP

Leu

Ser

STOP

Tr P

Leu

Pro

His

Arg

Leu

Pro

His

Arg

Leu

Pro

Gin

Arg

Leu

Pro

Gin

Arg

lie

Thr

Asn

Ser

lie

Thr

Asn

Ser

lie

Thr

Lys

Arg

Met

Thr

Lys

Arg

Val

Ala

Asp

Gly

Val

Ala

Asp

Gly

Val

Ala

Glu

Gly

Val

Ala

Glu

Gly

One- and three-letter abbreviations for amino acids

Ala

Alanine

Asx

Asparagine or aspartate

Cys

Cysteine

Asp

Aspartate

Glu

Glutamate

Phe

Phenylalanine

Gly

Glycine

His

Histidine

lie

Isoleucine

Lys

Lysine

Leu

Leucine

Met

Methionine

Asn

Asparagine

Pro

Proline

Gin

Glutamine

Arg

Arginine

Ser

Serine

Thr

Threonine

Val

Valine

Trp

Tryptophan

Tyr

Tyrosine

Glx

Glutamate or glutamine

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